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 &amp;deg;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...

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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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Research

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

Isolation of Brain and Spinal Cord Mononuclear Cells Using Percoll Gradients

Isolation of immune cells that infiltrate the central nervous system (CNS) during infection, trauma, autoimmunity or neurodegeneration, is often required to define their phenotype and effector functions. Histochemical approaches are instrumental to determine the location of the infiltrating cells and to analyze the associated CNS pathology. However, in-situ histochemistry and immunofluorescent staining techniques are limited by the number of antibodies that can be used at a single time to characterize immune cell subtypes in a particular tissue. Therefore, histological approaches in conjunction with immune-phenotyping by flow cytometry are critical to fully characterize the composition of local CNS infiltration. This protocol is based on the separation of CNS cellular suspensions over discontinous percoll gradients. The current article describes a rapid protocol to efficiently isolate mononuclear cells from brain and spinal cord tissues that can be effectively utilized for identification of various immune cell populations in a single sample by flow cytometry.

The current article describes a rapid protocol to efficiently isolate mononuclear cells from brain and spinal cord tissues that can be effectively utilized for flow cytometric analyses.

Isolation of immune cells that infiltrate the central nervous system (CNS) during infection, trauma, autoimmunity or neurodegeneration, is often required ...

Immunology and Infection

Quantitative Assessment of Immune Cells in the Injured Spinal Cord Tissue by Flow Cytometry:  a Novel Use for a Cell Purification Method

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true quantitative analysis of cellular inflammation. Flow cytometry is a quick alternative method to quantify immune cells in the injured brain or spinal cord tissue. Historically, flow cytometry has been used to quantify immune cells collected from blood or dissociated spleen or thymus, and only a few studies have attempted to quantify immune cells in the injured spinal cord by flow cytometry using fresh dissociated cord tissue. However, the dissociated spinal cord tissue is concentrated with myelin debris that can be mistaken for cells and reduce cell count reliability obtained by the flow cytometer. We have advanced a cell preparation method using the OptiPrep gradient system to effectively separate lipid/myelin debris from cells, providing sensitive and reliable quantifications of cellular inflammation in the injured spinal cord by flow cytometry. As described in our recent study (Beck &amp; Nguyen et al., Brain. 2010 Feb; 133 (Pt 2): 433-47), the OptiPrep cell preparation had increased sensitivity to detect cellular inflammation in the injured spinal cord, with counts of specific cell types correlating with injury severity. Critically, novel usage of this method provided the first characterization of acute and chronic cellular inflammation after SCI to include a complete time course for polymorphonuclear leukocytes (PMNs, neutrophils), macrophages/microglia, and T-cells over a period ranging from 2 hours to 180 days post-injury (dpi), identifying a surprising novel second phase of cellular inflammation. Thorough characterization of cellular inflammation using this method may provide a better understanding of neuroinflammation in the injured CNS, and reveal an important multiphasic component of neuroinflammation that may be critical for the design and implementation of rational therapeutic treatment strategies, including both cell-based and pharmacological interventions for SCI.

Quantification of cellular inflammation in the injured/pathological CNS by flow cytometry is complicated by lipid/myelin debris that can have similar size and granulation to cells, decreasing sensitivity/accuracy. We have advanced a cell preparation method to remove myelin debris and improve cell detection by flow cytometry in the injured spinal cord.

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true ...

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Isolation and Flow Cytometric Analysis of Glioma-infiltrating Peripheral Blood Mononuclear Cells

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 malignant gliomas soon after intracranial engraftment if the cancer cells are rendered deficient in their expression of the &#946;-galactoside-binding lectin galectin-1 (gal-1). More recent work now shows that a population of Gr-1+/CD11b+ myeloid cells is critical to this effect. To better understand the mechanisms by which NK and myeloid cells cooperate to confer gal-1-deficient tumor rejection we have developed a comprehensive protocol for the isolation and analysis of glioma-infiltrating peripheral blood mononuclear cells (PBMC). The method is demonstrated here by comparing PBMC infiltration into the tumor microenvironment of gal-1-expressing GL26 gliomas with those rendered gal-1-deficient via shRNA knockdown. The protocol begins with a description of&#160;how to culture and prepare GL26 cells for inoculation into the syngeneic C57BL/6J mouse brain. It then explains the steps involved in the isolation and flow cytometric analysis of glioma-infiltrating PBMCs from the early brain tumor microenvironment. The method is adaptable to a number of in vivo experimental designs in which temporal data on immune infiltration into the brain is required. The method is sensitive and highly reproducible, as glioma-infiltrating PBMCs can be isolated from intracranial tumors as soon as 24 hr post-tumor engraftment with similar cell counts observed from time point matched tumors throughout independent experiments. A single experimentalist can perform the method from brain harvesting to flow cytometric analysis of glioma-infiltrating PBMCs in roughly 4-6 hr depending on the number of samples to be analyzed. Alternative glioma models and/or cell-specific detection antibodies may also be used at the experimentalists&#8217; discretion to assess the infiltration of several other immune cell types of interest without the need for alterations to the overall procedure.

Presented here is a straightforward method for the isolation and flow cytometric analysis of glioma-infiltrating peripheral blood mononuclear cells that yields time-dependent quantitative data on the number and activation status of immune cells entering the early brain tumor microenvironment.

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 ...

Flow Cytometry Protocols for Surface and Intracellular Antigen Analyses of Neural Cell Types

Flow cytometry has been extensively used to define cell populations in immunology, hematology and oncology. Here, we provide a detailed description of protocols for flow cytometric analysis of the cluster of differentiation (CD) surface antigens and intracellular antigens in neural cell types. Our step-by-step description of the methodological procedures include: the harvesting of neural in vitro cultures, an optional carboxyfluorescein succinimidyl ester (CFSE)-labeling step, followed by surface antigen staining with conjugated CD antibodies (e.g., CD24, CD54), and subsequent intracellar antigen detection via primary/secondary antibodies or fluorescently labeled Fab fragments (Zenon labeling). The video demonstrates the most critical steps. Moreover, principles of experimental planning, the inclusion of critical controls, and fundamentals of flow cytometric analysis (identification of target population and exclusion of debris; gating strategy; compensation for spectral overlap) are briefly explained in order to enable neurobiologists with limited prior knowledge or specific training in flow cytometry to assess its utility and to better exploit this powerful methodology.

We provide a detailed description of a protocol for flow cytometric analysis of surface antigens and/or intracellular antigens in neural cell types. Critical aspects of experimental planning, step-by-step methodological procedures, and fundamental principles of flow cytometry are explained in order to enable neurobiologists to exploit this powerful technology.

Flow cytometry has been extensively used to define cell populations in immunology, hematology and oncology. Here, we provide a detailed description of ...

Analysis of Microglia and Monocyte-derived Macrophages from the Central Nervous System by Flow Cytometry

Numerous studies have demonstrated the role of immune cells, in particular macrophages, in central nervous system (CNS) pathologies. There are two main macrophage populations in the CNS: (i) the microglia, which are the resident macrophages of the CNS and are derived from yolk sac progenitors during embryogenesis, and (ii) the monocyte-derived macrophages (MDM), which can infiltrate the CNS during disease and are derived from bone marrow progenitors. The roles of each macrophage subpopulation differ depending on the pathology being studied. Furthermore, there is no consensus on the histological markers or the distinguishing criteria used for these macrophage subpopulations. However, the analysis of the expression profiles of the CD11b and CD45 markers by flow cytometry allows us to distinguish the microglia (CD11b+CD45med) from the MDM (CD11b+CD45high). In this protocol, we show that the density gradient centrifugation and the flow cytometry analysis can be used to characterize these CNS macrophage subpopulations, and to study several markers of interest expressed by these cells as we recently published. Thus, this technique can further our understanding of the role of macrophages in mouse models of neurological diseases and can also be used to evaluate drug effects on these cells.

This protocol provides an analysis of the macrophage subpopulations in the adult mouse central nervous system by flow cytometry and is helpful for the study of multiple markers expressed by these cells.

Numerous studies have demonstrated the role of immune cells, in particular macrophages, in central nervous system (CNS) pathologies. There are two main ...

High-Dimensionality Flow Cytometry for Immune Function Analysis of Dissected Implant Tissues

The success of implanting laboratory-grown tissue or a medical device in an individual is subject to the immune response of the recipient host. Considering an implant as a foreign body, a hostile and dysregulated immune response may result in the rejection of the implant, while a regulated response and regaining of homeostasis can lead to its acceptance. Analyzing the microenvironments of implants dissected out under in vivo or ex vivo&#160;settings can help in understanding the pattern of immune response, which can ultimately help in developing new generations of biomaterials. Flow cytometry is a well-known technique for characterizing immune cells and their subsets based on their cell surface markers. This review describes a protocol based on manual dicing, enzymatic digestion, and filtration through a cell strainer for the isolation of uniform cell suspensions from dissected implant tissue. Further, a multicolor flow cytometry staining protocol has been explained, along with steps for initial cytometer settings to characterize and quantify these isolated cells by flow cytometry.

Isolation of cells from dissected implants and their characterization by flow cytometry can significantly contribute to understanding the pattern of immune response against implants. This paper describes a precise method for the isolation of cells from dissected implants and their staining for flow cytometric analysis.

The success of implanting laboratory-grown tissue or a medical device in an individual is subject to the immune response of the recipient host. ...

Bioengineering

Flow Cytometric Analysis of Lymphocyte Infiltration in Central Nervous System during Experimental Autoimmune Encephalomyelitis

Multiple sclerosis (MS) is an autoimmune disease of the central nervous system (CNS) caused by the combination of environmental factors and susceptible genetic background. Experimental autoimmune encephalomyelitis (EAE) is a typical disease model of MS widely used for investigating the pathogenesis in which T lymphocytes specific for myelin antigens initiate an inflammatory reaction in CNS. It is very important to assess how lymphocytes in the CNS regulate the development of disease. However, the approach for measuring the quantity and quality of infiltrated lymphocytes in the CNS is very limited due to the difficulties in isolating and detecting infiltrated lymphocytes from the brain. This manuscript presents a protocol for that is useful for the isolation, identification, and characterization of infiltrated lymphocytes from the CNS and will be helpful for our understanding of how lymphocytes are involved in the development of the CNS autoimmune disease.

This manuscript presents a protocol to induce active experimental autoimmune encephalomyelitis (EAE) in mice. A method for the isolation and characterization of the infiltrated lymphocytes in the central nervous system (CNS) is also presented to show how lymphocytes are involved in the development of CNS autoimmune disease.

Multiple sclerosis (MS) is an autoimmune disease of the central nervous system (CNS) caused by the combination of environmental factors and susceptible ...

Flow Cytometry Analysis of Immune Cell Subsets within the Murine Spleen, Bone Marrow, Lymph Nodes and Synovial Tissue in an Osteoarthritis Model

Osteoarthritis (OA) is one of the most prevalent musculoskeletal diseases, affecting patients suffering from pain and physical limitations. Recent evidence indicates a potential inflammatory component of the disease, with both T-cells and monocytes/macrophages potentially associated with the pathogenesis of OA. Further studies postulated an important role for subsets of both inflammatory cell lineages, such as Th1, Th2, Th17, and T-regulatory lymphocytes, and M1, M2, and synovium-tissue-resident macrophages. However, the interaction between the local synovial and systemic inflammatory cellular response and the structural changes in the joint is unknown. To fully understand how T-cells and monocytes/macrophages contribute towards OA, it is important to be able to quantitively identify these cells and their subsets simultaneously in synovial tissue, secondary lymphatic organs and systemically (the spleen and bone marrow). Nowadays, the different inflammatory cell subsets can be identified by a combination of cell-surface markers making multi-color flow cytometry a powerful technique in investigating these cellular processes. In this protocol, we describe detailed steps regarding the harvest of synovial tissue and secondary lymphatic organs as well as generation of single cell suspensions. Furthermore, we present both an extracellular staining assay to identify monocytes/macrophages and their subsets as well as an extra- and intra-cellular staining assay to identify T-cells and their subsets within the murine spleen, bone marrow, lymph nodes and synovial tissue. Each step of this protocol was optimized and tested, resulting in a highly reproducible assay that can be utilized for other surgical and non-surgical OA mouse models.

Here, we describe a detailed and reproducible flow cytometry protocol to identify monocyte/macrophage and T-cell subsets using both extra- and intracellular staining assays within the murine spleen, bone marrow, lymph nodes and synovial tissue, utilizing an established surgical model of murine osteoarthritis.