The authors would like to acknowledge the support from NIH/NINDS R01 NS069861 and NJCBIR CBIR14IRG024 to VS.

1. Biocytin Filling during Electrophysiology
NOTE: The readers can refer to alternative sources for basic patch-clamp recording techniques and instrumentation19-22, which are not elaborated upon here. The steps detailed here assume that the equipment and procedures for patch-clamp recordings are already established, and the description will be restricted to details related to biocytin-filling and post-hoc immunostaining. All experiments outlined in this manuscript were performed on rats.
<ol>
	<li>Using a vibratome, prepare live brain sections of the desired region at a thickness of 300 - 350 &#181;m23.</li>
	<li>Prepare an internal solution containing 0.2% biocytin by adding 2 mg of biocytin to 1 mL of internal solution6. For best results, sonicate the internal solution for 10 - 15 min until the biocytin dissolves completely. Next, load the internal solution in a 1&#160;mL syringe fitted with a 0.2&#160;&#181;m pore size polypropylene filter tip attached to a microloader. Keep this loading syringe on a cold pack to maintain the stability of the Mg-ATP and Na-GTP in the internal solution. 
	NOTE: An internal solution containing potassium gluconate (126 mM K-gluconate, 4 mM KCl, 10 mM HEPES, 4 mM Mg-ATP, 0.3 mM Na-GTP, and 10 mM phosphocreatine) is optimal for the recovery of certain neurochemical markers, including parvalbumin, during immunolabeling. In our experience, using cesium- and chloride-based internal solutions reduces the ability to recover immunostaining for certain neurochemical markers. Neurobiotin or a cell-impermeant, fixable, polar tracer combining an Alexa Fluorophore with biocytin can be used as an alternative to biocytin.</li>
	<li>Under infrared differential interference contrast, visualize the appropriate cell for recording. To minimize any disruption of the axon, approach the cell body by lowering the pipette along its axis using the &#34;approach&#34; function available in most micromanipulators. Establish whole-cell recordings under infrared differential interference contrast using patch-clamp physiology protocols19-22. 
	NOTE: Avoid approaching the cell from the direction of the putative axon, as it will sever the axon, visualized as a bleb at the cut end (green arrow in Figure&#160;1). Also, avoid applying high positive pressure to the recording pipette while approaching the cell to minimize&#160;the spill of biocytin, which will result in high background staining.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54880/54880fig1.jpg" /> 
Figure 1. Suggested Plane for Approaching Cells for Biocytin Fills. The image illustrates a Granule Cell (GC) fill with biocytin during recording that has been processed to reveal biocytin (in red using 594-conjugated streptavidin). The GC has its dendrites oriented in the XY plane. Notice the severed granule cell axon (green arrow) due to pipette movement along the XY plane. Note that it is ideal to approach this neuron along the XZ plane. Scale bar: 50 &#181;m.
<ol start="4">
	<li>In voltage clamp configuration, determine the whole-cell capacitance and series resistance in response to small (5 mV), brief (30&#160;ms) voltage steps using the using the membrane test function in bath mode in an electrophysiology data acquisition and analysis software. This will ensure the establishment of whole-cell recording conditions to allow for biocytin-filling.</li>
	<li>Conduct physiological recordings in either current- or voltage-clamp mode as needed. A minimum recording time of &#62;10 min at 34 &#176;C is ideal. 
	NOTE: Longer recording times may be needed for studies performed at room temperature.</li>
	<li>Upon completion of the physiological recordings, re-establish the patch-clamp configuration by slowly moving the recording pipette in small steps, alternating upward (along the Z-axis) and outward (along the X-axis) in voltage-clamp mode.</li>
	<li>Simultaneously monitor the capacitance and input resistance using the &#34;membrane test&#34; function to visualize the loss of capacitive transients and the collapse of the current responses to a straight line, indicating the re-sealing of the cell and the establishment of an outside-out-patch at the pipette tip. Holding the cell at a depolarized potential (-40 mV) will facilitate the re-sealing process. 
	NOTE: This procedure for the removal of a cell typically ensures the complete recovery of the soma and dendrites. Do not apply positive pressure to the recording pipette during this procedure or while detaching the cell from the pipette. Visualization of the soma of the recorded cell in the slice indicates successful disengagement. The presence of cellular debris or a membrane patch at the end of the detached tip typically indicates dislocation of the soma and will not yield complete cellular morphology.</li>
	<li>After detaching the pipette from the cell, retain the slice in the recording chamber for 3 - 5 min to ensure the transport of the dye to distal dendritic and axonal processes.</li>
	<li>Transfer the slices to a 24-well plate containing 4% PFA&#160;for fixation. 
	CAUTION: PFA is carcinogenic, and appropriate personal protective equipment, including gloves and a face mask, must be used to avoid skin irritation and inhalation. PFA is flammable and must be kept out of reach of fire. PFA is never disposed of in the drain and must be collected as chemical waste. PFA fixation must be isolated from areas utilized for live slice preparation to avoid contamination of the physiology setup, including transfer pipettes.</li>
	<li>24 - 48 h after PFA fixation, transfer the slices to 0.1 M Phosphate-buffered Saline (PBS) for storage prior to immunostaining. 
	NOTE: Ideally, biocytin recovery and immunostaining are best when performed soon after fixation, although staining performed within 7 d&#160;of fixation yields good results. Slices can be maintained in PBS with 0.02 - 0.05% sodium azide for staining performed up to and over 90 days after fixation. Although overnight fixation in PFA works well for most antibodies, it is possible that some antibodies work best when the duration of incubation in fixative is reduced. Fixation in PFA for over 48 h reduces the availability of antigens and diminishes the chances of successful secondary immunostaining for neurochemical markers. 
	CAUTION: Sodium azide is extremely hazardous and an irritant upon contact with the skin or eyes or upon ingestion or inhalation. Severe over-exposure can result in death. The carcinogenicity and mutagenicity of the compound are not known. Use appropriate personal protective equipment including gloves, splash goggles, a lab coat, and a dust respirator. Sodium azide is never disposed of in the drain and must be collected as chemical waste.</li>
</ol>
2. Staining of the First Primary Antibody and Biocytin
(Day 1) 
NOTE: The following immunostaining procedure is for free-floating sections and requires continuous shaking at a low speed (2 revolutions/min, rpm) on a shaker for all incubation steps.
<ol>
	<li>Wash the tissue 3x for 10 min each in 0.1 M PBS (pH = 7.4). 
	NOTE: 0.1 M PBS can be commercially purchased or made in the laboratory as follows (makes 1 L): 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 in 1 L of dH20. 
	NOTE: If the desired neurochemical marker is known, staining for a protein/peptide can be performed simultaneously with biocytin staining (steps 2.2 - 2.4). If staining for biocytin only, proceed to step 2.5.</li>
	<li>OPTIONAL: Block with 10% Blocking Serum (BS; normal goat serum (NGS) diluted in 0.3% Triton X-100 in PBS for 1 h at RT). 
	NOTE: Each well should receive the same volume of the solution; the recommended volume is between 250 - 500 &#181;L/well. The selection of the blocking serum is based on the species in which the secondary antibodies are raised (e.g., NGS is used for dye-conjugated goat anti-(respective primary antibody species)). If the need arises to use secondary antibodies raised in different species to immunostain sections, include 10% of each normal serum in blocking step (2) and 3% of each normal serum for the primary and secondary antibody incubation steps.</li>
	<li>(OPTIONAL) Incubate the sections in primary antibody, CB1R (polyclonal, guinea pig) with 0.3% Triton X-100, and 3% BS in 0.1 M PBS at RT O/N. CB1R is used to identify cannabinoid receptor type 1 expression. 
	NOTE: This step is optional and not required to reveal the biocytin filling. Figure&#160;2 illustrates a section labeled for biocytin and CB1R during the first staining process. O/N incubation at RT is sufficient for most of the primary antibodies; however, certain primary antibodies, including CB1R, require 3 - 5 d of incubation. If the incubation needs to be longer than 1 d, it is recommended to incubate at 4 &#176;C because Triton X-100 can permeabilize slices and can lead to the disintegration of the tissue during prolonged incubation at room temperature. Include a negative control lacking primary antibody in at least one section for each series of experiments as a control for the specificity of the secondary antibody. For negative controls, the primary antibody is omitted, but the 0.3% Triton X-100 in 0.1 M PBS and BS are added.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54880/54880fig2.jpg" /> 
Figure 2. Successful CCK Staining 1 Week Following the Recovery of Biocytin Staining and Cannabinoid Receptor Type 1 (CB1R)-labeling. (A - C) Confocal images at 60X showing the biocytin-filled neuron (A) and CB1R immunoreactivity (B), indicated by the arrowhead. The same section was processed for CCK immunostaining after 1 week (C). The overlay of images is shown in D. CCK immunoreactivity was revealed using far-red and was pseudo-colored in cyan. (E) Morphological reconstruction of the biocytin-filled cell in A. Note that the biocytin and CB1R staining are evident even after the second CCK immunostaining. Also note the expected distribution of CB1R and CCK immunostaining patterns. (F - H) Magnified image of the axon from the cell, as in A, show close co-localization of biocytin and CB1R in axons (arrow heads). Scale bar: 50 &#181;m (A - D), 100 &#181;m (E), and 10 &#181;m (F - H). Images are reproduced from Yu et al. 20159. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54880/54880fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(Day 2)
<ol start="4">
	<li>Wash the brain sections 3x for 10 min each in 0.1 M PBS (pH = 7.4).</li>
	<li>Incubate the sections in streptavidin red dye conjugate (concentration: 1:1,000; excitation: 594 nm with emission in the visible red spectrum) with 0.3% Triton X-100 and 3% BS in 0.1 M PBS at 4 &#176;C O/N in the dark (wrapped in aluminum foil) to reveal the biocytin. OPTIONAL: Include a secondary antibody, goat anti-guinea pig (concentration: 1:500; excitation: 488 nm and emission in green), with the above incubation if following the optional primary incubation in step 2.3. 
	NOTE: Secondary antibody will be needed only if the optional primary antibody staining (step 2.3) is performed. To visualize biocytin, a streptavidin-fluorophore conjugate with an emission spectrum in red is recommended, as it helps to visualize finer axons in detail and with better contrast (Figures&#160;1 - 3). During double or triple immunostaining, in order to avoid cross-reactivity of secondary antibodies with each other, add secondary antibodies one after the other in a serial fashion (2&#160;h interval between serial additions of the antibodies is sufficient). Figure&#160;3A illustrates a biocytin-filled cell processed without the optional primary antibody staining and imaged prior to resectioning.</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/54880/54880fig3.jpg" /> 
Figure 3. Successful Second Immunostaining following the Recovery of Biocytin Morphology and Resectioning. (A1) Confocal image of a 300&#160;&#181;m slice showing the recovery of the dendritic morphology of a biocytin-filled cell. (A2) Membrane voltage traces of the neuron in response to +500 and -100 pA current injections. (B) Recovery of the biocytin-filled soma in a 50&#160;&#181;m section obtained after resectioning the 300&#160;&#181;m section (in A1) after mounting and imaging. (B2) Subsequent immunostaining of the thin section revealed colocalization of the biocytin-filled soma (right panel) with CCK (middle panel). The merged image is illustrated in the right panel. Scale bar: 50 &#181;m. Panel A2 is reproduced with permission from Yu et al., in press25. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54880/54880fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(Day 3)
<ol start="6">
	<li>Wash brain sections 3x for 10 min each in 0.1 M PBS (pH = 7.4).</li>
	<li>To protect the fluorescence and to facilitate re-staining in the future, mount the sections in an aqueous-based mounting medium and seal the edges of the coverslip with clear nail polish. 
	NOTE: It is best to restain sections as early as possible to avoid difficulties in removing the nail polish from the slide, mold infestation, and dehydration of tissue due to leakage of the mounting medium from the glass slide. Microbial contamination can be prevented by using 0.02% sodium azide in PBS. 
	CAUTION: Sodium azide is highly toxic and flammable when it is in contact with water. Please refer to standard operating procedures for the handling and disposal of hazardous chemicals. 
	(Days 4 - 7)</li>
	<li>Perform confocal imaging with excitation at 594 and 488 nm and at a magnification of 20X or 40X to reveal biocytin-filled neurons in red and neurochemical markers (CB1R) in green. Assess colabeling (Figure&#160;2).</li>
	<li>Set the camera exposure to less than 100 ms to avoid photobleaching and obtain confocal image stacks of the axonal and dendritic arbors of the entire neuron. Use confocal image stacks and neuron tracing software packages to reconstruct the neuronal morphology.</li>
</ol>
3. Staining of the Second Primary Antibody
NOTE: While the second immunostaining step can be performed 90 d&#160;after the first staining, optimal staining is achieved if the second primary antibody staining is performed within 7 - 10 d.
(After 7 - 90 d)
<ol>
	<li>Apply acetone to a cotton swab and strip the nail polish off of the glass slide.</li>
	<li>Remove the coverslip carefully using fine-tip forceps and put a few droplets of 0.1 M PBS on the sections.</li>
	<li>Carefully remove the sections from the slide using a fine paint brush and wash them in 0.1 M PBS for 24 - 48 h on a shaker covered in aluminum foil in low light at 4 oC. 
	NOTE: It is crucial to cover sections with aluminum foil to avoid photobleaching.</li>
	<li>To ensure complete removal of the mounting medium, wash the sections 1 - 2x&#160;on the following day in 0.1 M PBS for 10 min each.</li>
	<li>Proceed with incubation in the second primary antibody (e.g., cholecystokinin, a neuropeptide found in certain GABAergic interneurons (CCK; concentration: 1:1,000; mouse monoclonal antibody)) for 1 d&#160;(Figure&#160;2). 
	NOTE: This procedure is similar to step 2.3 but uses a different antibody. Additionally, the blocking step is omitted during re-staining.</li>
	<li>Wash the brain sections three times for 10 min each in 0.1 M PBS (pH = 7.4).</li>
	<li>Stain with the fluorophore-conjugated secondary antibody goat anti-mouse (concentration: 1:500; excitation wavelength: 647 nm), making sure that the excitation-emission spectra of the secondary antibody do not overlap with streptavidin (concentration: 1:1,000; excitation wavelength: 594 nm) and prior immunofluorescence stains.</li>
	<li>Mount the sections in aqueous mounting medium and seal the edges of the cover slip with clear nail polish. 
	NOTE: Store the sections at 4 &#176;C in an opaque storage box to protect the fluorescence.</li>
</ol>

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The authors have nothing to disclose.

Critical Steps within the Protocol 
Filling the patched cell with biocytin is the most crucial step to ensure the full recovery of the morphology. For full recovery of the cell, it is essential to select an optimal slice orientation to minimize the severing of processes during slicing. This orientation may differ based on the circuit and cell type under examination. Next, it is essential to allow adequate time for the biocytin to diffuse to the dendrites and axons. Some dyes, such as neurobiotin, can also penetrate neurons coupled to the recorded cell through gap junctions. The size of the pipette tip also needs to be optimized, as the soma tends to be pulled out when the opening is large and biocytin loading is compromised when the pipette tip is too small. Additionally, appropriate procedures while disengaging from whole-cell mode are essential to recover the soma. After recording for 24 h, the subsequent incubation of the slices in 4% PFA followed by PBS ensures that the cross-linking of the tissues is reduced, which is critical for immunohistochemistry. Storage of the slices at 4 &#176;C is also crucial for maintaining slice integrity. However, prolonged storage in PFA reduces eventual success with antibody-labeling procedures. Optimizing the concentration and duration of the primary antibody incubation is important, as these will differ for individual antibodies, and some antibodies may require from 3 - 5 d&#160;of incubation for optimal penetration in thick tissue. To ensure that the slices are undamaged before the secondary primary antibody staining, adequate care is needed during the removal of the cover slip.
Modifications and Troubleshooting
We have observed that there is no set duration between the first and second stain. Sections have been re-stained months later on the same tissue and with more than one stain. In our recent works, we compared sections processed using the re-staining method with those using standard immunostaining protocols and find no obvious difference in the staining patterns or the robustness of biocytin fills6,9,25,26.
While it remains to be tested, it is possible that re-staining may be performed more than once, with tissue integrity being a limiting factor for repeated handling. The availability of discrete fluorophores also poses a limit for the number of antigens that can be visualized. Therefore, careful handling of the tissue is advised. An additional process that remains to be tested is restaining using antigen recovery protocols (for surface receptors). Since the integrity of the tissue is retained after removal from the cover slip, we expect that the procedure should also work in staining protocols that recommend antigen-recovery. In this regard, physiological recordings often yield cells with no known neurochemical markers6,9. The procedures detailed here allow for the recovery of their morphology based on biocytin immunostaining. While the neurochemical identity may not be known a priori, re-staining for potential markers could facilitate identification of markers for neurons that have been described on the basis of morphology and electrical properties.
Limitations of the Technique
A limitation associated with performing immunohistochemical staining on thick sections is the insufficient penetration of the antibody through the complete thickness of the section, especially with standard overnight incubation in the primary antibody. Moreover, different antibodies and streptavidin, used to reveal biocytin, can have differential tissue penetration. Thus, it is recommended that trial runs are performed to assess the depth of antibody penetration and to modify the incubation times, temperatures, and antibody and Triton&#160;X-100 concentrations in order to achieve optimal slice penetration and staining for each antibody. It should also be noted that successful labeling with one antibody or with streptavidin-biocytin does not imply that a second antibody will penetrate to the same depth of the section. Therefore, care must be taken to verify the penetration of each antibody to the level at which the biocytin-labeled cell processes are examined for co-localization with the neurochemical marker. If the penetration is inadequate, the slice can be re-sectioned at &#60;60 &#181;m for immunostaining. Note, however, that since the slices were already processed for biocytin, the morphology of the cell can be captured by confocal imaging to eliminate the possible loss of anatomical data during re-sectioning and to facilitate neuronal reconstruction. While it is possible that the age and animal species may alter the degree of antibody penetration, we have successfully used these procedures in rat tissue from 30-day- and over 60-day-old rats and in mice.
A second limitation is that biocytin is not inherently fluorescent and cannot be used for live imaging and real-time morphological characterization during recording. Alternative unconjugated fluorophores and Lucifer yellow have been utilized for live imaging of the cells27. They are not permanent and lose fluorescence upon fixation, making it impractical for post-hoc neurochemical identification of the recorded neuron. Recently, fluorophores with biocytin have been introduced to overcome this limitation and can be used for live imaging as well as for post-hoc processing in double- or triple-labeling, as detailed here. Interestingly, to date, biocytin fills have resulted in better electrical and staining properties for combined electrophysiological and anatomical studies when compared to Lucifer yellow28.
Significance of the Technique with Respect to Existing/Alternative Methods
The major advantages of using the protocol detailed here are that, unlike resectioning, the biocytin-filled cell structure and processes remain intact and do not lead to the permanent loss of tissue, nor to a need for laborious reconstructions from multiple sections. Thus, a single-cell fill followed by immunocytochemistry can help in the morphological reconstruction and identification of specific neurochemical markers.
Future Applications or Directions after Mastering This Technique
In all, we present a novel practical approach for the recovery of morphology and post-hoc double- and triple-immunolabeling of neurons following electrophysiological recordings. The process is particularly advantageous for finding new markers as knowledge evolves, re-evaluating neurons that were previously filled, visualized, and imaged. It is particularly advantageous because the immunohistochemistry can be performed weeks or even months later, saving tissue and antibodies. The method does not require any special chemicals and can safely be performed in any laboratory.

The brain is known for diversity in the structural and functional characteristics of its individual neuronal elements. Understanding the roles of distinct neuronal types in brain function and pathology requires characterization and unambiguous identification of neurons. Structurally, the morphological features defined by somato-dendritic location determine the potential inputs that a given neuron receives, while the pattern of axonal arborization identifies potential postsynaptic targets. The structural diversity of neurons has been appreciated since the days of Ram&#243;n y Cajal&#39;s seminal histological studies1. The advent of single-cell recording techniques revealed that structurally distinct neurons also show differences in firing patterns and synaptic characteristics. The diversity in structure and physiology is particularly evident in GABAergic inhibitory neurons2,3. In addition, it has become increasingly apparent that structurally similar neurons can express different neurochemical markers and show corresponding functional differences4. Similarly, neurons with the same neurochemical markers can have distinct structures and functions5-10. Thus, in practice, the analysis of the functional characteristics of neurons and their role in the network entails defining both the morphological and neurochemical identities. Even with the advent of reporter mouse lines targeting specific neurochemical markers, it is often necessary to determine morphology and subtype identity based on immunohistology11.
The standard method used to characterize cells recorded in acute brain slices is to fill them with biocytin or neurobiotin during the recording, fix the sections in paraformaldehyde (PFA) following the recordings, and use immunohistochemistry to reveal the morphology and neurochemistry. Since the thickness of sections for slice physiology are typically 300 &#181;m or more, and because most antibodies fail to penetrate all the way through that depth, the slices need to be re-sectioned to 60 &#181;m or less to allow for simultaneous immunostaining for biocytin and neurochemical markers12-14. Unfortunately, resectioning is laborious; risks loss of tissue during sectioning; and can lead to differential tissue shrinkage, complicating morphological reconstructions. Additionally, prior knowledge of morphology could help narrow down the candidate markers that are likely to be expressed by the cells. We have modified the standard biocytin immunohistology protocols to allow serial processing of sections first for the recovery of morphology and then for the identification of potential neurochemical markers.
Immunohistochemistry is the study of antigen distribution in tissues or cells and can be visualized using an enzyme, fluorescent labels, radioactive elements, or gold colloid particles15. The procedure involves using primary antibodies to specifically tag and amplify one or more specific antigens, followed by the use of fluorescent secondary antibodies targeting the primary antibody for visualization. Due to the need to distinguish the fluorescence spectra of each secondary antibody without overlap, only a limited number of antigens can be examined simultaneously. Thus, prior knowledge of morphology could be useful in selecting the candidate neurochemical markers for cell classification. Conceptually, the rationale behind serial processing of already-stained sections is based on the premise that immunolabeling for one protein or peptide should not interfere with antigenicity and subsequent immunolabeling for a structurally independent peptide16. This lack of interference is due to the binding of the antibodies to a specific protein epitope on an antigen and therefore allows for the simultaneous staining of multiple antigens in the same tissue. The number of antigens revealed by immunostaining is limited by the need for non-overlapping spectra of the fluorescent secondary antibodies and by the need to target individual antigens with antibodies raised in different species so as to eliminate cross-reactivity17,18. While this is the reasoning behind serial rather than simultaneous labeling with two distinct antibodies that may interact, to our knowledge, immunostaining for a second antigen has not been reported after the completion of immunolabeling for one or more antigens on mounted sections. Here, we describe a method for serial immunostaining of previously stained and mounted sections. While we detail this process for a serial immunolabeling procedure for the recovery of morphology followed by staining for protein/peptide markers in thick sections, the same procedures can be used in standard, thin histological sections as well. In addition, we describe a practical approach to fill recorded neurons with biocytin and the process to dislodge the electrode from the cell upon completion of recordings to optimize the filling of the axonal and dendritic arbors of neurons, as presented in our recent work6,8.
The most crucial advantage of the procedure described here is that the morphology of the recorded cell can be fully recovered and imaged before attempting to resection or immunostain the slices. Although issues with the penetration of certain antibodies may render it necessary to resection slices for secondary immunostaining, the procedures detailed here would eliminate the need to reconstruct complex neurons from multiple sections and would avoid issues due to tissue loss and differential shrinkage, which can compromise reconstruction following resectioning. An added advantage is that the process will reduce cost, time, effort, and expensive antibodies by limiting immunostaining and re-sectioning to slices in which biocytin-filled neurons are recovered. The most practical aspect is the additional immunostaining that can be performed on sections stained months before using the aforementioned technique. In particular, the recovery of morphology would considerably reduce the potential that physiological data from the cells is discarded due to an inability to obtain a basic morphological characterization of the cell type.

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immunostaining of biocytin-filled and processed sections for neurochemical markers

Electrophysiological recordings of cells using the patch clamp technique have allowed for the identification of different neuronal types based on firing patterns. The inclusion of biocytin/neurobiotin in the recording electrode permits post-hoc recovery of morphological details, which are necessary to determine the dendritic arborization and the regions targeted by the axons of the recorded neurons. However, given the presence of morphologically similar neurons with distinct neurochemical identities and functions, immunohistochemical staining for cell-type-specific proteins is essential to definitively identify neurons. To maintain network connectivity, brain sections for physiological recordings are prepared at a thickness of 300 &#181;m or greater. However, this thickness often hinders immunohistological postprocessing due to issues with antibody penetration, necessitating the resectioning of the tissue. Resectioning of slices is a challenging art, often resulting in the loss of tissue and morphology of the cells from which electrophysiological data was obtained, rendering the data unusable. Since recovery of morphology would limit data loss and guide in the selection of neuronal markers, we have adopted a strategy of recovering cell morphology first, followed by secondary immunostaining. We introduce a practical approach to biocytin filling during physiological recordings and subsequent serial immunostaining for the recovery of morphology, followed by the restaining of sections to determine the neurochemical identity. We report that sections that were filled with biocytin, fixed with paraformaldehyde (PFA), stained, and coverslipped can be removed and restained with a second primary antibody days later. This restaining involves the removal of the coverslip, the washing of sections in a buffer solution, and the incubation of primary and secondary antibodies to reveal the neurochemical identity. The method is advantageous for eliminating data loss due to an inability to recover morphology and for narrowing down the neurochemical markers to be tested based on morphology.

Upon successful completion, the sections retain the biocytin fill and the immunolabeling performed in step 2 and can be imaged using confocal or epifluorescence microscopy. In addition, the processed sections will also show immunostaining for the antigen labeled during the subsequent processing in step 3. In the section illustrated in Figure&#160;2, the morphology of a biocytin-filled neuron in a thick section (300 &#181;m) was revealed using streptavidin visualized in red and the first primary (CB1R) visualized in green. The first immunostaining was performed within 7 d&#160;of PFA fixation. The slice was mounted in an aqueous mounting medium, imaged using a confocal microscope, and stored for a period of two months before re-staining for the CCK antibody. Note the morphology of the recorded neuron, including axonal arbors (Figure&#160;2E), was reconstructed from the confocal images obtained after the first staining procedure. Colocalization of the axon with CB1R immunoreactivity (Figure&#160;2 F - H) was expected based upon physiological studies. The second staining for CCK was performed on sections two months after the first staining procedure. CCK was imaged in far-red and was pseudo-colored in cyan for visualization. Similarly, Figure&#160;3 illustrates an example of a cell in which biocytin-labeling was revealed and imaged in a thick section (Figure&#160;3A1). The intrinsic physiology of the neuron is illustrated in Figure&#160;3A2. The slice was re-sectioned at 50 &#181;m and, although some sections containing dendritic processes were lost, the section containing the soma recovered (Figure&#160;3B1). Subsequent CCK immunostaining of the thin section showed CCK expression in green in the biocytin-labeled soma. Mounting the thick slice in an aqueous medium prevents excessive tissue shrinkage. On average, 300-&#181;m sections mounted using the aqueous medium shrink by about 25.67 &#177; 3.14% (n = 5 slices) when measured one week after mounting. Thus, the thick sections mounted using the aqueous medium are ~225 &#181;m, allowing for resectioning. In several control studies, it was confirmed that the process of immunolabeling for the second time did not lead to non-specific labeling by the antibody. For example, it is known that parvalbumin (PV) and CCK immunoreactivity are mutually exclusive in hippocampal neurons. Controls in which the first immunolabeling step (2) for biocytin (red) and CCK (green) was followed by the second labeling for parvalbumin (with excitation at 405 nm and emission in blue) showed non-overlapping expression patterns (Figure&#160;4). Additionally, the distinctive laminar axonal staining patterns of the neurons expressing specific neurochemical markers (PV, CCK, CB1R) were not altered by the serial re-staining procedures (Figures&#160;2 - 4). Moreover, secondary staining consistently revealed co-labeling of the biocytin-filled neurons with the neurochemical markers expected on the basis of the physiological recordings. Thus, we are confident that the restaining procedure does not lead to loss of specificity in labeling.
<img alt="Figure 4" src="/files/ftp_upload/54880/54880fig4.jpg" /> 
Figure 4. Lack of Overlap between Mutually Exclusive Markers on the Second Immunostaining Following Prior Immunostaining and Mounting of Sections. (A - C) Confocal images at 10X showing a biocytin-filled neuron (A) in red, indicated by an arrowhead, and CCK immunoreactivity in green (B), indicated by the arrow. Note the lack of overlap. The same section was processed for parvalbumin (PV) immunostaining in blue after more than 2 months (C). The overlay of images is shown in D. Note that the PV axons are distributed in the granule cell layer, with CCK in the molecular layer in the expected nonoverlapping pattern. Also note the lack of colocalization of the two markers in neuronal somata. The zoomed images to the right illustrate that the biocytin-labeled neuron expresses PV (arrowhead) and not CCK (arrow). Scale bar: 100 &#181;m (A - D), 20 &#181;m (panels to the right). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54880/54880fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
One common reason for sub-optimal biocytin fills is the accidental severing of the neuronal processes with the recording pipette while approaching the cell. A lost appendage of the cell appears as a bleb of biocytin at the end of an axon, as illustrated in Figure&#160;1 (green arrow). Axonal blebs are often visualized when patching superficial cells, whose axons may be severed during slicing procedures. However, optimization of the plane of slicing and knowledge of the axonal distribution pattern of the targeted cell class can aid in avoiding approaching the soma from the direction of the putative axon, which can also sever the axon, resulting in the appearance of an axonal bleb. In most cell types, it is preferable to approach the cell body along the Z-axis (white arrow with mark). A second issue concerns the application of excessive positive pressure to the pipette while approaching the cell. This can not only damage the cell and its connections but also causes the internal solution containing biocytin to spill around the cell, leading to high background fluorescence. Background staining also increases when the time taken to approach the cell is prolonged or when slice quality is not optimal, which can compromise the ability to define cellular morphology (Figure&#160;5A). It is possible to pull the soma out while dislodging the pipette. This will result in a lack of somatic morphology (Figure&#160;5B). To avoid pulling out the cell, move the pipette up and out in small steps using the &#34;fine&#34; movement settings of the manipulator. If the cell appears to be attached to the pipette through a long membrane extension, tap/flick the manipulator gently to dislodge the pipette from the cell. Avoid using the coarse manipulator settings until the cell has fully dislodged.
<img alt="Figure 5" src="/files/ftp_upload/54880/54880fig5.jpg" /> 
Figure 5. Images Illustrate Unsuccessful Biocytin Fills. (A) Image showing high background fluorescence due to excessive positive pressure in the recording pipette, resulting in poor cell visibility. Finer structures of neuronal processes, including axons, are visualized beyond the region of the spill. Also, notice that soma has been pulled out. (B) A neuron with the soma pulled out during pipette detachment. Arrows show successful fills of the axon and dendrites, whereas the soma and appendages proximal to the arrows have not been recovered. (C) A cell patched in a superficial layer of the slice (&#60;50 &#181;m from surface) results in filled cells exhibiting cut axon and dendrites (arrowheads). Notice the beading (arrows) of granule cell dendrites, indicating poor cell health (C). Scale bar: 50 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54880/54880fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Finally, in order to recover complete cellular morphology without severed axons or dendrites, it is best to target cell &#62;50 &#181;m deep to the slice surface. Superficial cells may have processes that have been cut and may lack the normal complement of synaptic inputs.
The most common pitfalls in the restaining procedures are associated with the removal of the coverslip and the recovery of the section without damaging the tissue. A second issue concerns antibody penetration into thick tissue, especially when incubated overnight. In several instances, better penetration has been achieved by incubating in the primary antibody for up to 72 h at 4 &#176;C on an orbital shaker placed in a cold room. Although re-sectioning the tissue is the most effective procedure to ensure antibody penetration, prolonging the duration of incubation in primary antibody or increasing the detergent concentration to 0.5 - 1% Triton-X100 may also aid antibody penetration24. Since most recorded neurons are located within 50 - 100 &#181;m from the surface of the section, we find that the staining of thick sections is adequate to label biocytin-filled somata or processes at this level. However, it is recommended that the extent of antibody labeling is checked on each section before interpreting negative co-localization results.

Watch this Scientific Journal Video about Immunostaining of Biocytin-filled and Processed Sections for Neurochemical Markers at JoVE.com

Immunostaining of Biocytin-filled and Processed Sections for Neurochemical Markers

This protocol presents a method for the morphological recovery of neurons patched during electrophysiological recordings using biocytin filling and subsequent immunohistochemical postprocessing. We show that thick biocytin-filled sections that were stained and coverslipped can be restained with a second primary antibody days&#160;or months later.

Electrophysiological recordings of cells using the patch clamp technique have allowed for the identification of different neuronal types based on firing ...

immunostaining-biocytin-filled-processed-sections-for-neurochemical

Nanyang Technological University

Research

JoVE Journal

Neuroscience

22.9K Views.  Rutgers New Jersey Medical School. This protocol presents a method for the morphological recovery of neurons patched during electrophysiological recordings using biocytin filling and subsequent immunohistochemical postprocessing. We show that thick biocytin-filled sections that were stained and coverslipped can be restained with a second primary antibody days or months later.

Video: Immunostaining of Biocytin-filled and Processed Sections for Neurochemical Markers

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Morphometric Analyses of Retinal Sections

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the retinal ganglion cell layer (RGCL) in rat models with N-methyl-D-aspartate (NMDA)&#x2013;induced excitotoxicity1, retinal ischemia-reperfusion injury2 and glaucoma3. Reduction of INL and inner plexiform layer (IPL) thicknesses were reversed with citicoline treatment in rats' eyes subjected to kainic acid-mediated glutamate excitotoxicity4. Alteration of RGC density and soma sizes were observed with different drug treatments in eyes with elevated intraocular pressure3,5,6. Therefore, having objective methods of analyzing the retinal morphometries may be of great significance in evaluating retinal pathologies and the effectiveness of therapeutic strategies.
The retinal structure is multi-layers and several different kinds of neurons exist in the retina. The morphometric parameters of retina such as cell number, cell size and thickness of different layers are more complex than the cell culture system. Early on, these parameters can be detected using other commercial imaging software. The values are normally of relative value, and changing to the precise value may need further accurate calculation. Also, the tracing of the cell size and morphology may not be accurate and sensitive enough for statistic analysis, especially in the chronic glaucoma model. The measurements used in this protocol provided a more precise and easy way. And the absolute length of the line and size of the cell can be reported directly and easy to be copied to other files. For example, we traced the margin of the inner and outer most nuclei in the INL and formed a line then using the software to draw a 90 degree angle to measure the thickness. While without the help of the software, the line maybe oblique and the changing of retinal thickness may not be repeatable among individual observers. In addition, the number and density of RGCs can also be quantified. This protocol successfully decreases the variability in quantitating features of the retina, increases the sensitivity in detecting minimal changes.
 	This video will demonstrate three types of morphometric analyses of the retinal sections. They include measuring the INL thickness, quantifying the number of RGCs and measuring the sizes of RGCs in absolute value. These three analyses are carried out with Stereo Investigator (MBF Bioscience &mdash; MicroBrightField, Inc.). The technique can offer a simple but scientific platform for morphometric analyses.

This video demonstrates three types of morphometric analyses of the retina, which include measuring the inner nuclear layer thickness, quantifying the number of retinal ganglion cells (RGCs) and measuring the sizes of RGCs. The technique can offer a simple but scientific platform for morphometric analyses.

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

Quadruple Immunostaining of the Olfactory Bulb for Visualization of Olfactory Sensory Axon Molecular Identity Codes

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory Sensory Neuron (OSN) is a bipolar cell with a single dendrite and a single unbranched axon. An OSN expresses only one Olfactory Receptor (OR) gene, OSNs expressing a given type of OR converge their axons to a few sets of invariant glomeruli in the Olfactory Bulb (OB). A remarkable feature of OSN projection is that the expressed ORs play instructive roles in axonal projection. ORs regulate the expression of multiple axon-sorting molecules and generate the combinatorial molecular code of axon-sorting molecules at the OSN axon termini. Thus, to understand the molecular mechanisms of OR-specific axon guidance mechanisms, it is vital to characterize their expression profiles at the OSN axon termini within the same glomerulus. The aim of this article was to introduce methods for collecting as many glomeruli as possible on a single OB section and for performing immunostaining using multiple antibodies. This would allow the comparison and analysis of the expression patterns of axon-sorting molecules without staining variation between OB sections.

Olfactory sensory neurons express a wide variety of axon-sorting molecules to establish proper neural circuitry. This protocol describes an immunohistochemical staining method to visualize combinatorial expressions of axon-sorting molecules at the axon termini of olfactory sensory neurons.

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory ...

Toluidine Blue Staining of Resin-Embedded Sections for Evaluation of Peripheral Nerve Morphology

Peripheral nerves extend throughout the body, innervating target tissues with motor or sensory axons. Due to widespread distribution, peripheral nerves are frequently damaged because of trauma or disease. As methods and strategies have been developed to assess peripheral nerve injury in animal models, function and regeneration, analyzing the morphometry of the peripheral nerve has become an essential terminal outcome measurement. Toluidine blue staining of nerve cross sections obtained from resin embedded nerve sections is a reproducible method for qualitative and quantitative assessments of peripheral nerves, enabling visualization of morphology number of axons and degree of myelination. This technique, as with many other histological methods, can be difficult to learn and master using standard written protocols. The intent of this publication is therefore to accentuate written protocols for toluidine blue staining of peripheral nerves with videography of the method, using sciatic nerves harvested from rats. In this protocol, we describe in vivo peripheral nerve fixation and collection of the tissue, and post-fixation with 2% osmium tetroxide, embedding of nerves in epoxy resin, and ultramicrotome sectioning of nerves to 1-2&#956;m thickness. Nerve sections then transferred to a glass slide and stained with toluidine blue, after which they are quantitatively and qualitatively assessed. Examples of the most common problems are shown, as well as steps for mitigating these issues.

Here we present a protocol to visualize fine structures of peripheral nerves by obtaining and staining 1-2 &#181;m sections with toluidine blue

Peripheral nerves extend throughout the body, innervating target tissues with motor or sensory axons. Due to widespread distribution, peripheral nerves ...

Identifying Cell Surface Markers of Primary Neural Stem and Progenitor Cells by Metabolic Labeling of Sialoglycan

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized niches in vivo and can be isolated and expanded in vitro, serving as an important resource for cell transplantation to repair brain damage. However, NSPCs are heterogeneous and not clearly defined at the molecular level or purified due to a lack of specific cell surface markers. The protocol presented, which has been previously reported, combines a neural-endothelial co-culture system with a metabolic glycan labeling method to identify the surface sialoglycoproteome of primary NSPCs. The NSPC-endothelial co-culture system allows self-renewal and expansion of primary NSPCs in vitro, generating a sufficient number of NSPCs. Sialoglycans in cultured NSPCs are labeled using an unnatural sialic acid metabolic reporter with bioorthogonal functional groups. By comparing the sialoglycoproteome from self-renewing NSPCs expanded in an endothelial co-culture with differentiating neural culture, we identify a list of membrane proteins that are enriched in NSPCs. In detail, the protocol involves: 1) set-up of an NSPC-endothelial co-culture and NSPC differentiating culture; 2) labeling with azidosugar per-O-acetylated N-azidoacetylmannosamine (Ac4ManNAz); and 3) biotin conjugation to modified sialoglycan for imaging after fixation of neural culture or protein extraction from neural culture for mass spectrometry analysis. Then, the NSPC-enriched surface marker candidates are selected by comparative analysis of mass spectrometry data from both the expanded NSPC and differentiated neural cultures. This protocol is highly sensitive for identifying membrane proteins of low abundance in the starting materials, and it can be applied to marker discovery in other systems with appropriate modifications

Presented here is a protocol that combines an in vitro neural-endothelial co-culture system and metabolic incorporation of sialoglycan with bioorthogonal functional groups to expand primary neural stem and progenitor cells and label their surface sialoglycoproteins for imaging or mass-spectrometry analysis of cell surface markers.

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized ...

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.

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 ...