This study was supported by the Deutsche Forschungsgemeinschaft (DFG; SFB1118). The authors would like to thank Axel Erhardt for performing most of the dissection and helpfully transmitting this knowledge, and Dr. Volker Eckstein for assisting in the technical aspect of the flow cytometer.

Wild-type C57BL/6 mice (Males; 10-12 weeks) were kept in standard 12 h light/dark cycle and were provided free access to standard chow diet and water. All animal experiments were conducted in accordance to the relevant guidelines by the local Animal Care and Use Committee and approved by the local Animal Care and Use Committee at the regional authority in Karlsruhe, Germany (G216/10).
1. Perfusion of the Mouse
<ol>
	<li>Sacrifice the mouse (C57BL/6; Males; 10-12weeks) using CO2 or in accordance with local regulations. 
	NOTE: Retro-orbital bleeding can be performed to obtain control leukocytes, using EDTA-coated 1.5 mL tubes. This method of bleeding is only efficient immediately after the sacrifice. Further, peritoneal cavity lavage (PCL) can be performed to obtain control macrophages, using pre-heated PBS at 37 &#176;C to avoid cold agglutination of the blood, which may hamper perfusion.</li>
	<li>Disinfect the mouse by spraying the fur on the abdomen liberally with 70% v/v ethanol and repeatedly wiping with a tissue with cotton gauze. Using blunt scissors, cut open the abdominal skin by a longitudinal incision.</li>
	<li>Expose the pleural cavity by first making an incision into the diaphragm using curved, blunt scissors and then extending the incision along the entire length of the rib cage. Make a similar cut on the contralateral side until the sternum can be carefully lifted away.</li>
	<li>With a clear view of the heart and the major vessels, insert a butterfly 23&#190; gauge needle, connected to a 50 mL syringe filled with ice-cold PBS, into the left chamber of the heart.</li>
	<li>Make an incision to the right atrium of the heart using iris scissors to create as large of an outlet as possible without damaging the descending aorta, and then gently depress the syringe to push through approximately 30 mL of PBS. The exudate should be running clear. The clearing of the liver and/or kidneys can be used as an indicator of a good perfusion.</li>
</ol>
2. Dissection of Sciatic Nerve and DRG
NOTE: For the dissection of the sciatic nerves, the skin above the gluteus maximus is removed and the mouse is positioned ventral side down whilst stretching out the lower limbs.
<ol>
	<li>Bilaterally dissect the whole sciatic nerves by separating the muscle tissue from the upper dorsal thigh, and follow the nerve up along the spinal column. Cut the nerve where it exits from the spinal column and immediately above the knee at the other end. 
	NOTE: Avoid excessive handling of the nerve such as crushing or pulling with forceps.</li>
	<li>Using forceps, transfer the excised nerve(s), approximately 2 cm per nerve, to a 1.5 mL tube containing 1 mL of ice-cold PBS and store on ice. 
	NOTE: The nerve can be stored overnight on ice. 
	NOTE: For the DRG dissection, using forceps and a scalpel, detach the skin from the underlying soft tissue by gently peeling off the remaining back skin to expose the cervical, thoracic, and lumbar spinal regions. Remove the head by cutting at the base of the skull using a blunt pair of scissors.</li>
	<li>Cut the ribs parallel with and close to the spinal column on both sides, before detaching the viscera connected to the anterior side of the spinal column. Cut the femurs and remove the spinal column by a transverse cut at the level of the femurs. Using iris scissors, cut away any muscle, fat, and other soft tissues from the spine.</li>
	<li>Place the spinal column with the dorsal side facing up and open it, starting at the caudal end, by sliding one tip of the scissors into the vertebral canal at the three o&#39;clock position and the bone snipped. Repeat the procedure at the nine o&#39;clock position. 
	NOTE: It is important to cut exactly in the center to avoid damaging the DRG.</li>
	<li>Repeat the cutting from side-to-side until the rostral end is reached and the spinal cord is fully exposed. Using a dissection microscope, remove the spinal cord to expose the spinal nerves, which can be used to locate the DRGs within the spine.</li>
	<li>Remove the lumbar DRGs by gently tugging on the dorsal root to pull the DRG into the vertebral canals, then clipping the nerves and connective tissue with iris scissors to remove the DRGs. 
	NOTE: It is possible to restrict the collection to the L4-L6 DRG, which are contributing to the sciatic nerve.</li>
	<li>Using forceps, transfer the DRGs to a 1.5 mL tube containing approximately 1 mL of ice-cold PBS, and store on ice. 
	NOTE: The nerve can be stored overnight on ice.</li>
</ol>
3. Digestion of Sciatic Nerve and DRG
<ol>
	<li>Prepare the digestion media (DMEM, 10 mM HEPES, 5 mg/mL BSA, 1.6 mg/mL collagenase Type 4) supplemented with 100 &#181;g/mL of deoxyribonuclease, and store on ice. 
	NOTE: The amount of digestion medium required per mouse (sciatic nerves + DRGs) is 1 mL. Once prepared, aliquots of the digestion media can be stored at -20 &#176;C.</li>
	<li>Transfer the sciatic nerves and DRGs from one mouse into separate 1.5 mL tubes containing 500 &#181;L of digestion media. For the sciatic nerves, chop the material into 15-20 pieces using spring scissors and then incubate the tissue suspensions in a shaking incubator at 37 &#176;C for 20 min with gentle agitation (&#8804; 450 rpm).</li>
	<li>Repeatedly triturate the cell suspension through 1,000 &#181;L pipette tip to mechanically dissociate any incompletely digested tissue. 
	NOTE: If the suspension is difficult to titrate, then cut the pipette tip with scissors to create a blunt end. Incubate the tissue suspension at 37 &#176;C for 20 min with gentle agitation (&#8804; 450 rpm).</li>
	<li>Repeat the trituration through a 1,000 &#181;L pipette tip and incubate for a further 10 min at 37 &#176;C with gentle agitation (&#8804; 450 rpm).</li>
	<li>Prepare the FACS buffer (PBS supplemented with 10% FCS and 1 mM EDTA) and store on ice. Rinse and soak a stainless-steel screen with a mesh size of 140 &#181;m with FACS buffer in 60 x 15 mm Petri dish on ice. 
	NOTE: Nylon screen filters should be avoided as it leads to a significant loss in the yield of nucleated cells.</li>
	<li>Repeatedly pass the tissue suspensions through the 140 &#181;m screen filter, as it is held over the Petri dish, with blunt forceps, on ice. Collect the cell suspension from the Petri dish and transfer it to a new 1.5 mL tube stored on ice.</li>
	<li>Push any incompletely digested tissue that remains on the filter through by repeated trituration with a 1,000 &#181;L pipette tip. Rinse the screen filter twice with 500 &#181;L of FACS buffer and collect the resulting cell suspension in the Petri dish to combine with the cell suspension from step 3.6. 
	NOTE: If the flow cytometer used is sensitive to debris, the cell suspension from step 3.7 can be passed through a 70 &#181;m screen filter; however, this may lead to a loss in the yield of nucleated cells.</li>
	<li>Centrifuge the cell suspensions at 300 x g for 5 min at 4 &#176;C. Discard the supernatant and resuspend the cell pellet in 1 mL of FACS buffer. Repeat this washing step once.</li>
	<li>Resuspend the cell pellet in 150-350 &#181;L of FACS buffer, depending upon the number of staining that are required (150 &#181;L FACS buffer per staining panel).</li>
</ol>
4. Staining of Sciatic Nerve and DRG with Fluorescently Labeled Antibodies for Flow Cytometry
<ol>
	<li>Transfer 100 &#181;L of the sciatic nerve or DRG cell suspension into a V-shaped well in a 96-well plate on ice. 
	NOTE: Single-stained cells are necessary for all antibodies used as they will provide the necessary controls for setting up the flow cytometry with respect to photomultiplier tube (PMT) voltage gains and compensation, as well as to help distinguish specific from non-specific binding. Due to the limited material available from the sciatic nerve and DRGs, blood leukocytes and macrophages from the PCL (steps 1.1-1.2) can be used as the single-stained controls, the staining for which should be performed in parallel with the staining of the sciatic nerve and DRGs.</li>
	<li>Pool the remaining cells suspensions (approximately 50 &#181;L per sample) from either the sciatic nerves or DRG samples, to serve as an unstained control.</li>
	<li>Centrifuge the plate at 400 x g for 5 min at 4 &#176;C. Discard the supernatant by inverting the plate over a bin and applying a sharp flick of the wrist, and gently tap dry on paper towels.</li>
	<li>For surface staining, resuspend each of the cell pellets in 20 &#181;L of FACS buffer containing the antibodies of interest (CD45-A647, CD11b-PerCP/Cy5.5, and MHC Class II-Biotin) and incubate, protected from light, on ice for 30 min. 
	NOTE: (1) The optimal antibody dilution should be determined prior to use by performing a dilution series according to the manufacturer&#39;s product information; (2) Non-specific stainings can be reduced by incubation with Fc-block at this step, according to the manufacturer&#39;s instructions; and (3) Isotype control stainings are generally not required when using primary antibodies directly conjugated with a fluorophore and/or when the antigen(s) of interest produce distinct populations.</li>
	<li>Add 130 &#181;L of FACS buffer to each well and centrifuge the plate at 400 x g for 5 min at 4 &#176;C. Discard the supernatant as previously described (step 4.3) and wash the cell pellets twice with 150 &#181;L of FACS buffer. 
	NOTE: If using an unconjugated primary antibody or a primary antibody conjugated to biotin, then an appropriate secondary reagent, conjugated to a fluorophore, can be added at this point and steps 4.4-4.5 are repeated. In the case of this protocol, streptavidin-PE/Cy7 or Streptavidin-PerCP was used to detect MHC Class II in combination with CD45-A647/CD11b-PerCP/Cy5.5, or in combination with CD68-APC and F4/80-PE/Cy7, respectively.</li>
	<li>For fixation, after the last washing step, resuspend the cell pellets in 100 &#181;L of 4% paraformaldehyde in PBS (pH 7.4). Incubate the plate on ice for 10 min and then centrifuge at 400 x g for 5 min at 4 &#176;C. Discard the supernatant as previously described (step 4.3) and wash the cell pellets once with 150 &#181;L of FACS buffer. 
	Caution: Paraformaldehyde is toxic; wear appropriate protection such as gloves and eyeglasses, etc. 
	NOTE: Resuspend the cell pellets in 150 &#181;L of FACS buffer and store protected from the light, at 4&#176; C for up to two days. If no intracellular staining is required, then proceed to step 4.11 after centrifuging the plate at 400 x g for 5 min at 4 &#176;C.</li>
	<li>For intracellular staining, resuspend the cell pellets in 150 &#181;L of FACS buffer+0.5% detergent, and incubate at room temperature, protected from the light for 15 min. Centrifuge the plate at 400 x g for 5 min at 4 &#176;C. Discard the supernatant as previously described (step 4.3) and wash the cell pellet once with 150 &#181;L of FACS buffer+0.5 % detergent. Resuspend the cell pellet in 20 &#181;L of FACS buffer+0.5% detergent containing the antibodies of interest (CD68-APC, CD206-PE) and incubate, protected from light, at room temperature for 30 min.</li>
	<li>Add 130 &#181;L of FACS buffer+0.5% detergent to each well and centrifuge the plate at 400 x g for 5 min at 4 &#176;C. Discard the supernatant as previously described (step 4.3) and wash the cell pellets twice with 150 &#181;L of FACS buffer+0.5 % detergent. 
	NOTE: If using an unconjugated primary antibody or a primary antibody conjugated to biotin, then an appropriate secondary reagent, conjugated to a fluorophore, can be added at this point and steps 4.9-4.10 repeated.</li>
	<li>Resuspend the cell pellets in 150 &#181;L of FACS buffer+0.5 % detergent and store, protected from the light, at 4 &#176;C for up to two days.</li>
	<li>Resuspend the cell pellets with 150 &#181;L FACS buffer+0.5 % detergent supplemented with 5 &#181;g/mL DAPI and incubate for 5 min at room temperature, protected from the light. Centrifuge the plate at 400 x g for 5 min at 4 &#176;C. Discard the supernatant as previously described (step 4.3).</li>
	<li>Resuspend the cell pellets in 150 &#181;L of PBS and transfer to an appropriately labeled FACs tube. Store at room temperature, protected from the light, until ready for analysis. 
	NOTE: After resuspension in PBS, the cells should be analyzed within hours, since the DAPI will slowly dissociate from the DNA, leading to a loss in signal.</li>
</ol>
5. Setup of Flow Cytometry and Running of Samples
<ol>
	<li>Ensure that the flow cytometer is equipped with the appropriate lasers and/or filter sets to measure the fluorophores used to stain the cells. This can be determined under the default cytometer configuration on the system. In the case of this protocol, cells were analyzed using a flow cytometer equipped with three lasers (405 nm, 488 nm, and 633 nm) allowing for the detection of &#8804; 5 fluorophores.</li>
	<li>Select the appropriate channels for detection of the fluorophores of interest. All fluorescent signals are best detected with logarithmic amplification.</li>
	<li>Select an appropriate flow rate for the samples. This parameter can vary depending upon the system used and should be determined empirically. In the case of this protocol, a flow rate of MED-to-HI (35-60 &#181;L/min) was used to prevent clogging of the needle.</li>
	<li>Select the total number of events/cells to be collected per sample. The minimum number of events should be 1,000,000.</li>
	<li>Create scatter plots under a global worksheet of the SSC-A versus FSC-A, as well for each fluorophore of interest versus FCS-A. Create a statistical view that will display the number of events and the mean fluorescent intensity (MFI) for the selected fluorophores.</li>
	<li>Set the voltage of the FSC-A and SSC-A channels, such that the smallest cells fit into the lower left 10% of the scatter plot. Determine the background fluorescence and minimum sample fluorescence from the unstained controls from either the sciatic nerves or DRGs. Determine the parent (P1) gate based upon the scatter plot from DAPI-only staining controls. 
	NOTE: This is not a collection gate, due in part, to the heterogeneous nature of the DAPI+ population in the PNS.</li>
	<li>Using the FSC scatter plots for each of the fluorophores of interest, adjust the voltages so that the cells of the unstained control fall within the lower quarter of their respective scatter plots. Gates are then created above this region to define those cells that are positive for each of the respective markers/fluorophores. The placement of a gate is confirmed by running of the single-stained control (step 4.1). 
	NOTE: Compensation for overlapping emission spectra can be performed at this point or applied retrospectively with the appropriate analysis software.</li>
	<li>Once the flow cytometer has been set up, as described, the samples can be run.</li>
	<li>After the run, the remaining sample material can be discarded and the FCS files can be exported for further analysis.</li>
</ol>

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The authors have no conflicts of interest in regard to this study.

Sciatic nerves contain a large proportion of lipids, such as cholesterol, due to the content of myelin around the axons. Since the properties of lipids change with temperature, different results may be obtained at different temperatures. To ensure cell preservation, all steps in this protocol after the digestion were performed on ice. Whilst consistency is recommended for the sake of reproducibility of the results, it may be possible to increase yield by performing the steps 3.6 and 3.7 at room temperature. If the nerves...

Immune cells which enter the PNS from the circulation, as defined by the expression of CD45 and CD11b, help to maintain the integrity of the nerve and play a role both in regeneration and degeneration1. Macrophages (defined by their expression of CD68 in mouse) can be skewed towards an inflammatory phenotype, expressing more MHC class II and CD86 on their surface (M1), or towards an anti-inflammatory phenotype, expressing more intracellular CD206 (M2)2. Skewing of the macrophage phenotype is a dynamic process regulated through Akt signaling3, reflecting the different tasks of macrophages in the defense against pathogens (M1) and the role in tissue regeneration (M2). Regeneration of an injured nerve first requires phagocytosis of myelin debris by macrophages in the nerve4,5, and anti-inflammatory (CD68+CD206+) M2 macrophages have been shown to promote axon outgrowth in the PNS6. Reduced recruitment or macrophages to the PNS, or impaired capacity for phagocytosis may result in impaired regeneration and maintenance of nerve integrity. Inflammatory M1 macrophages, expressing MHC class II, are less capable of phagocytosis than M2 macrophages, and neuronal inflammation is implicated in the pathogenesis of several neurodegenerative diseases7.
The changes that occur to the nerve resident immune system as a consequence of damage may be quantitative, manifesting in loss of CD45+ leukocytes (or increased infiltration of CD45+ leukocytes in the case inflammation), or qualitative, such as change of macrophage phenotype from M2 to M1 phenotype. The immune cells of the PNS have traditionally been analyzed by means of IF, using frozen sections or paraffin-embedded material8. IF is required for determining localization of the cells of interest. However, quantification using IF often relies on counting a relatively small number of cells in a narrow section of the tissue, making quantification unreliable and vulnerable to selection bias. For identification of specific subsets of immune cells, the simultaneous detection of extracellular and/or intracellular markers is required, whilst determination of the macrophage phenotype requires at least at least two markers, specifically CD206 and MHC class II. As most commonly available microscopes are limited to at least two-color channels, such as fluorescein isothiocyanate (FITC) and phycoerythrin (PE), the characterization of the specific subsets of immune cells by IF can be restrictive and incomplete, requiring the need to have multiple slides, derived from the same area of interest, which are stained and analyzed in parallel. This time-consuming aspect therefore does not necessary lend itself to the analysis of large sample sets. Furthermore, as most of the markers of interest are extracellular, the detection in tissue, which has either been embedded in paraffin or cryoconserved, can be problematic due to the disruption of membrane integrity and the masking of epitopes, as well as the loss of the antigens of interest from the use of solvents, such as acetone and methanol9.
In contrast, flow cytometry, which measures optical and fluorescence characteristics of single cells in suspension as they pass through a beam of light, provides a more practical and comprehensive means for analysis of the cell populations. Flow cytometry, rather than producing a digital image of the tissue, provides an automated quantification of set parameters, which include a cell&#39;s relative size and reflective index, referred to as the forward scatter (FSC), granularity/internal complexity or side-scatter (SSC), and relative fluorescence intensity, providing that the cell has been labeled with an appropriate fluorophore, such as a conjugated antibody. A typical flow cytometer consists of two, air-cooled lasers; an argon laser produces blue light at 488 nm and a helium-neon laser produces light at 633 nm. This combination allows for the detection and measurement, simultaneously, of at least five targets either on the surface or within the intracellular compartment. More advanced flow cytometers can consist of multiple lasers, which allow for the detection of up to eight different fluorochromes at once, providing that the peak emission wavelengths of the selected fluorochromes do not overlap significantly.
For analyzing by flow cytometry, the tissue of interest must first be enzymatically digested, generally with collagenase, to generate a single-cell suspension. The analysis of murine sciatic nerves has previously been difficult due to the small amount of tissue obtained from each mouse. In addition, the high fat content of the myelin around the axons hampers cell recovery and produces large amounts of debris. The method described herein for sciatic nerve preparation and digestion was adapted from the Schwann cell isolation protocol of Barrette et al.7, and aims to isolate enough cells from nerves of individual mice for flow cytometry analysis, in order to reduce variation between mice. Using DAPI for the identification of single cells in the raw tissue digest circumvents the need to remove axon debris, which commonly leads to cell loss. Washing several times with a detergent-rich buffer aids in the release of cells trapped within the fatty debris, thereby increasing the yield. Digestion of both full-length sciatic nerves from a single mouse according to this protocol generates &#8805;30,000 single nucleated events and at least 3 times that number was retrieved from the DRG. The proportion of CD45+ leukocytes was approximately 5% of total cell content in the sciatic nerve digest and approximately 5-10% in the DRG digest. The majority of the CD45+ cells in the sciatic nerve expressed the macrophage markers, CD68 and CD206.

<table>
	<tbody>
		<tr>
			<td>C57BL/6 Mouse</td>
			<td>Charles River</td>
			<td>C57BL/6NCrl</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM (+1g/L glucose, Glutamine, Pyruvate)</td>
			<td>Thermo</td>
			<td>31885023</td>
			<td>The source of this material is not important</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type 4</td>
			<td>Worthington Biochemical Corp., US</td>
			<td>LS004188</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease (DNase) I from bovine pancreas I</td>
			<td>Sigma-Aldrich</td>
			<td>DN25-1g</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Sigma-Aldrich</td>
			<td>E6758</td>
			<td></td>
		</tr>
		<tr>
			<td>Foetal Calf Serum (FCS), heat inactivated</td>
			<td>Sigma-Aldrich</td>
			<td>F4135</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody against CD11b-PerCP/Cy5.5</td>
			<td>Biolegend</td>
			<td>101228</td>
			<td>Clone M1/70</td>
		</tr>
		<tr>
			<td>Antibody against MHC class II, biotinylated</td>
			<td>Biolegend</td>
			<td>107603</td>
			<td>Clone M5/114.15.2</td>
		</tr>
		<tr>
			<td>Antibody against CD45-A647</td>
			<td>Biolegend</td>
			<td>107603</td>
			<td>Clone 30-F11</td>
		</tr>
		<tr>
			<td>Antibody against CD68-APC</td>
			<td>Biolegend</td>
			<td>137007</td>
			<td>Clone FA/11</td>
		</tr>
		<tr>
			<td>Antibody against CD206-PE</td>
			<td>Biolegend</td>
			<td>141706</td>
			<td>Clone C068C2</td>
		</tr>
		<tr>
			<td>Antibody against F4/80-PE/Cy7</td>
			<td>Biolegend</td>
			<td>123113</td>
			<td>Clone BM8</td>
		</tr>
		<tr>
			<td>Streptavidin-PE/Cy7</td>
			<td>Biolegend</td>
			<td>405206</td>
			<td>Used to detect MHCII-biotin with CD45-A647 and CDllb-PerCP/Cy5.5</td>
		</tr>
		<tr>
			<td>Streptavidin-PerCP</td>
			<td>Biolegend</td>
			<td>405213</td>
			<td>Used to detect MHCII-biotin with CD68-APC and F4/80-PE/Cy7</td>
		</tr>
		<tr>
			<td>Triton-X 100</td>
			<td>Sigma-Aldrich</td>
			<td>T8787</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma-Aldrich</td>
			<td>D9542-1MG</td>
			<td>Dilute in PBS to a 50x and store at 4 &#176;C in the dark</td>
		</tr>
		<tr>
			<td>Cell dissociation sieve - tissue grinder kit</td>
			<td>Sigma-Aldrich</td>
			<td>CD1</td>
			<td></td>
		</tr>
		<tr>
			<td>V-Shaped, 96-well plates</td>
			<td>Greiner/Sigma</td>
			<td>M8185</td>
			<td></td>
		</tr>
		<tr>
			<td>5ml polystryene round-bottom tube, 12x75mm</td>
			<td>BD Biosciences</td>
			<td>352008</td>
			<td></td>
		</tr>
		<tr>
			<td>60x15mm petri dish</td>
			<td>Greiner/Sigma</td>
			<td>Z643084</td>
			<td></td>
		</tr>
		<tr>
			<td>BD LSR II Flow Cytometer</td>
			<td>BD Biosciences</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

analysis of immune cells in single sciatic nerves and dorsal root ganglion from a single mouse using flow cytometry

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of the PNS are affected by injury and disease, affecting the nerve function and the capacity for regeneration. Neuronal immune cells are commonly analyzed by immunofluorescence (IF). While IF is essential for determining the location of the immune cells in the nerve, IF is only semi-quantitative and the method is limited to the number of markers that can be analyzed simultaneously and the degree of surface expression. In this study, flow cytometry was used for quantitative analysis of leukocyte infiltration into sciatic nerves or dorsal root ganglions (DRGs) of individual mice. Single cell analysis was performed using DAPI and several proteins were analyzed simultaneously for either surface or intracellular expression. Both sciatic nerves from one mouse that were treated according to this protocol generated &#8805; 30,000 single nucleated events. The proportion of leukocytes in the sciatic nerves, determined by expression of CD45, was approximately 5% of total cell content in the sciatic nerve and approximately 5-10% in the DRG. Although this protocol focuses primarily on the immune cell population within the PNS, the flexibility of flow cytometry to measure a number of markers simultaneously means that the other cells populations present within the nerve, such as Schwann cells, pericytes, fibroblasts, and endothelial cells, can also be analyzed using this method. This method therefore provides a new means for studying systemic effects on the PNS, such as neurotoxicology and genetic models of neuropathy or in chronic diseases, such as diabetes.

Cell suspensions from both full-length sciatic nerves and all major DRG were prepared, according to the protocol, from six healthy C57BL/6 mice and divided into three equal aliquots for staining. Counter staining with DAPI, which stains DNA, allows for the detection of a single cell population (Figure 1A-B, upper panel). Washing out the DAPI, prior to analysis by flow cytometry, decreased to an extent the fluorescent intensit...

Watch this Scientific Journal Video about Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry at JoVE.com

Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry

Quantitative analysis of cell content within the murine sciatic nerve is difficult due to the scarcity of the tissue. This protocol describes a method for tissue digestion and preparation that provides sufficient cells for flow cytometry analysis of immune cell populations from nerves of individual mice.

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of ...

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JoVE Journal

Neuroscience

11.4K Views.  University Hospital of Heidelberg. Quantitative analysis of cell content within the murine sciatic nerve is difficult due to the scarcity of the tissue. This protocol describes a method for tissue digestion and preparation that provides sufficient cells for flow cytometry analysis of immune cell populations from nerves of individual mice.

Video: Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

A Single-fly Assay for Foraging Behavior in Drosophila

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster. In a light-tight box illuminated by red light that is invisible to fruit flies, a camera linked to custom data acquisition software monitors the position of six flies simultaneously. Each fly is confined to walk in individual arenas containing a food odor at the center. The testing arenas rest on a porous floor that functions to prevent odor accumulation. Latency to locate the odor source, a metric that reflects olfactory sensitivity under different physiological states, is determined by software analysis. Here, we discuss the critical mechanics of running this behavioral paradigm and cover specific issues regarding fly loading, odor contamination, assay temperature, data quality, and statistical analysis.

A Single-fly Assay for Foraging Behavior in Drosophila

In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster.

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...

An Approach to Enhance Alignment and Myelination of Dorsal Root Ganglion Neurons

Axon regeneration is a chaotic process due largely to unorganized axon alignment. Therefore, in order for a sufficient number of regenerated axons to bridge the lesion site, properly organized axonal alignment is required. Since demyelination after nerve injury strongly impairs the conductive capacity of surviving axons, remyelination is critical for successful functioning of regenerated nerves. Previously, we demonstrated that mesenchymal stem cells (MSCs) aligned on a pre-stretch induced anisotropic surface because the cells can sense a larger effective stiffness in the stretched direction than in the perpendicular direction. We also showed that an anisotropic surface arising from a mechanical pre-stretched surface similarly affects alignment, as well as growth and myelination of axons. Here, we provide a detailed protocol for preparing a pre-stretched anisotropic surface, the isolation and culture of dorsal root ganglion (DRG) neurons on a pre-stretched surface, and show the myelination behavior of a co-culture of DRG neurons with Schwann cells (SCs) on a pre-stretched surface.

This protocol describes the isolation of dorsal root ganglion (DRG) neurons isolated from rats and the culture of DRG neurons on a static pre-stretched cell culture system to enhance axon alignment, with subsequent co-culture of Schwann Cells (SCs) to promote myelination.

Axon regeneration is a chaotic process due largely to unorganized axon alignment. Therefore, in order for a sufficient number of regenerated axons to ...

Single-cell RNA-Seq of Defined Subsets of Retinal Ganglion Cells

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for the improved understanding of neuronal diversity, it is important to identify genes whose expression defines various subpopulations of cells. The retina serves as an excellent model for the study of central nervous system diversity, as it is composed of multiple major cell types. The study of each major class of cells has yielded genetic markers that facilitate the identification of these populations. However, multiple subtypes of cells exist within each of these major retinal cell classes, and few of these subtypes have known genetic markers, although many have been characterized by morphology or function. A knowledge of genetic markers for individual retinal subtypes would allow for the study and mapping of brain targets related to specific visual functions and may also lend insight into the gene networks that maintain cellular diversity. Current avenues used to identify the genetic markers of subtypes possess drawbacks, such as the classification of cell types following sequencing. This presents a challenge for data analysis and requires rigorous validation methods to ensure that clusters contain cells of the same function. We propose a technique for identifying the morphology and functionality of a cell prior to isolation and sequencing, which will allow for the easier identification of subtype-specific markers. This technique may be extended to non-neuronal cell types, as well as to rare populations of cells with minor variations. This protocol yields excellent-quality data, as many of the libraries have provided read depths greater than 20 million reads for single cells. This methodology overcomes many of the hurdles presented by Single-cell RNA-Seq and may be suitable for researchers aiming to profile cell types in a straightforward and highly efficient manner.

Here, we present a combinatorial approach for classifying neuronal cell types prior to isolation and for the subsequent characterization of single-cell transcriptomes. This protocol optimizes the preparation of samples for successful RNA Sequencing (RNA-Seq) and describes a methodology designed specifically for the enhanced understanding of cellular diversity.

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for ...

Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each other anatomically, it is important to identify an appropriate model to use for the study of axon regeneration. By using a suitable model, we can formulate a specific treatment based on the severity of injury, the neuronal cell type of interest, and the desired spinal tract for assessing regeneration. Within the sensory pathway, DRG neurons are responsible for relaying sensory information from the periphery to the CNS. We present here a protocol that uses a DRG injection with a viral vector and a concurrent dorsal root crush injury in the lower cervical spinal cord of an adult rat as a model to study sensory axon regeneration. As demonstrated using a control virus, AAV5-GFP, we show the effectiveness of a direct DRG injection in transducing DRG neurons and tracing sensory axons into the spinal cord. We also show the effectiveness of the dorsal root crush injury in denervating the forepaw as an injury model for evaluating axon regeneration. Despite the requirement for specialized training to perform this invasive surgical procedure, the protocol is flexible, and potential users can modify many parts to accommodate their experimental requirements. Importantly, it can serve as a foundation for those in search of a suitable animal model for their studies. We believe that this article will help new users to learn the procedure in a very efficient and effective manner.

This protocol presents the use of a dorsal root ganglion (DRG) injection with a viral vector and a concurrent dorsal root crush injury in an adult rat as a model to study sensory axon regeneration. This model is suitable for investigating the use of gene therapy to promote sensory axon regeneration.

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each ...

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive targets for injection of novel therapeutics aimed at treating chronic pain. In small animal models, laminectomy has been used to facilitate DRG injection because it involves surgical removal of the vertebral bone surrounding each DRG. We demonstrate a technique for intraganglionic injection of lumbar DRG in a large animal species, namely, swine. Laminotomy is performed to allow direct access to DRG using standard neurosurgical techniques, instruments, and materials. Compared with more extensive bone removal via laminectomy, we implement laminotomy to conserve spinal anatomy while achieving sufficient DRG access. Intraoperative progress of DRG injection is monitored using a non-toxic dye. Following euthanasia on post-operative day 21, the success of injection is determined by histology for intraganglionic distribution of 4&#39;,6-diamidino-2-phenylindole (DAPI). We inject a biologically inactive solution to demonstrate the protocol. This method could be applied in future preclinical studies to target therapeutic solutions to DRG. Our methodology should facilitate testing the translatability of intraganglionic small animal paradigms in a large animal species. Additionally, this protocol may serve as a key resource for those planning preclinical studies of DRG injection in swine.

We describe a method for laminotomy in swine that provides access to lumbar dorsal root ganglia (DRG) for intraganglionic injection. Injection progress is monitored intraoperatively and histologically confirmed up to 21 days after surgery. This protocol could be used for future preclinical studies involving DRG injection.

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...