We would like to thank Andrew Lim, Ph.D. and Giles Best, Ph.D. for their help in setting up the flow cytometer. This project was supported by the Deutsche Forschungsgemeinschaft (DFG) (DFG-HA 8481/1-1) awarded to PH.

Northern Sydney Local Health District Animal Ethics Committee has approved all procedures mentioned in this protocol. Mice are housed and cared for in accordance with the Guide for the Care and Use of Laboratory Animals (National Health and Medical Research Council of Australia Revised 2010). For all experiments 10-12-week-old, male C57BL/6 mice were utilized.
NOTE: To induce post-traumatic OA, surgical destabilization of the medial meniscus (DMM) in the right stifle joint was performed. Detailed information regarding this animal model was published by Glasson et al.18. In short, general anesthesia is induced in an induction chamber using isoflurane and thereafter maintained using a nose cone. The surgical leg is shaved with a razor blade and the surgical site is washed and swabbed with ethanol to minimize contamination. The animal is then moved to the operating microscope and placed on a sterile towel and the leg draped with sterile paper drape to isolate the surgical site and minimize contamination. Using the microscope, a 0.5 cm medial para-patella arthrotomy is made, the patella luxated laterally, and the infra-patella fat pad elevated to expose the medial menisco-tibial ligament, which is transected with dissecting forceps. The joint is flushed with sterile saline to remove any blood and the wound is closed in three layers &#8211; joint capsule, subcutaneous tissue (using suture material) and skin (using surgical tissue glue). Methods described in this protocol, however, can be applied to other models and methods for inducing OA. OA can be induced in either side of the animal, and when harvesting tissues, it is important to harvest the ipsilateral (draining) lymph nodes.
1. Isolation of the spleen, contralateral bone marrow, ipsilateral lymph nodes draining the stifle and synovial tissue
<ol>
	<li>Euthanize the mouse by cervical dislocation. Place the mouse in a supine position under a dissecting microscope and wipe the chest, abdomen and legs with 70% ethanol. Carefully open the skin on the midline for the length of the abdomen using straight scissors leaving the abdominal cavity intact.</li>
	<li>Gently pull the skin on the right side of the animal away from the underlying muscle leaving the subcutaneous adipose tissue attached to the skin. Normally, gentle traction alone will separate the skin and underlying adipose tissue from the muscle. Sporadic adherences should be cut through with fine scissors in order to keep the tension necessary to separate tissues to a minimum and reduce the chance of harming the lymph nodes. Identify a crossing of three vessels by gently teasing out the adipose tissue located at the thigh using two curved fine forceps. The inguinal lymph node is located at the crossing and can be identified by its ovoid shape and slightly darker color.</li>
	<li>Remove the inguinal lymph node using fine dissecting forceps. Be careful not to rupture the capsule. Remove the remaining fat on the surface of the lymph node with the forceps.</li>
	<li>Open the abdominal cavity and identify the spleen. Cut out the spleen with fine scissors. Gently pull the intestines aside to expose the aorta and its bifurcation being careful not to harm them, in order to minimize risk of contamination.&#160;The iliac lymph node is located at the terminal segment of the abdominal aorta and the origin of the common iliac artery. Remove right iliac lymph node and proceed as described in 1.3.</li>
	<li>Gently remove the skin of both hind limbs. Dissect the left femur by cleaning it from muscle tissue using the blade and fine scissors. Carefully disconnect both the stifle and hip joint leaving the whole bone intact and remove the femur.</li>
	<li>Identify the patella tendon of the right stifle joint, then remove the adjoining muscle tissue proximal to this using fine scissors until approximately 5mm of quadriceps tendon proximal to the patella is exposed. Thereafter, cut through the quadriceps tendon approximately 3-4mm proximal to the patella to form a handle and using fine forceps gently pull it away from the joint. This will render the edges of the joint capsule attachment to femur visible, and using a scalpel blade carefully cut along the edges of the joint capsule on both sides, starting at the femur going towards the tibia, in order to maximize the amount of synovial membrane that is harvested. Whilst cutting it is important to maintain a gentle traction on the quadriceps tendon and pause when the synovial tissue block is only attached to the tibia. At this stage the intraarticular fat pad is now clearly visible distal to the patella and can be gently detached from the joint and anterior aspect of the menisci using the blade. Thereafter, cut along the remaining part of the joint capsule (tibial portion) to remove the synovial tissue block. 
	NOTE: This step has to be done very precisely to allow reliable results. After completing the dissection, the &#8220;synovial tissue block&#8221; should consist of the patella, patella tendon, infrapatellar fat pad, supra- and infrapatellar recesse synovial lining and associated joint capsule anterior to the collateral ligaments. Keep all tissues moist during dissection using 0.9% saline solution. 
	NOTE: Place each synovial tissue block sample in a separate well of a labelled 24-well plate containing 1.5 ml of RPMI 1640 medium. Combine both the iliac and the inguinal lymph node into one well and pool tissues from two mice.&#160;</li>
</ol>
2. Generation of single cell suspensions from each tissue
NOTE: In order to ensure sufficient cell numbers for flow analysis synovial tissues from two mice need to be pooled. In the current protocol, pool all tissues from the same two mice in order to maintain analogy. Furthermore, iliac and inguinal lymph nodes were combined for each animal resulting in a total of 4 lymph nodes for each sample. In general, cell numbers in spleen, bone marrow and lymph nodes from one animal are sufficient to conduct flow analysis and the protocol can be applied. However, when using tissues from only one animal lysing times might need to be adjusted.
<ol>
	<li>The spleen
	<ol>
		<li>Place the two pooled spleens onto a 70 &#181;m cell strainer on top of a 15 mL tube. Gently macerate the spleens through the mesh filter using a sterile 3 mL syringe plunger. Flush the strainer frequently with a total of 6 mL of RPMI 1640 medium supplemented with 10% FBS.</li>
		<li>Spin the cells (500 x g, 5 min, RT) and resuspend the pellet in 5 mL of red blood cell (RBC) lysis buffer. Incubate for 5 min at RT and stop the reaction by diluting the lysis buffer with 10 mL of PBS. Spin the cells (500 x g, 5 min, RT) and repeat this step once or until no more RBC are in the pellet. 
		NOTE: Refilter the suspension using a 30 &#181;m cell strainer into a new 15 mL tube between the two rounds of lysing to remove coagulated cells.</li>
		<li>After lysing is complete, spin the cells (500 x g, 5 min, RT), discard supernatant and resuspend pellet in 1 mL of PBS. Count the number of live cells on a hemocytometer using Trypan blue exclusion.</li>
	</ol>
	</li>
	<li>The lymph nodes
	<ol>
		<li>Place the four pooled lymph nodes onto a 70 &#181;m cell strainer on top of a 15 mL tube. Gently tease the lymph nodes apart into a single cell suspension by pressing with a sterile 3 mL syringe plunger. Flush the strainer frequently with a total of 6 mL of RPMI with 10% FBS.</li>
		<li>Spin the cells (500 x g, 5 min, RT), discard supernatant and resuspend the pellet in 500 &#181;L of PBS. Refilter the suspension using a 30 &#181;m cell strainer into a new 15 mL tube to remove coagulated cells. Count the number of live cells on a hemocytometer using Trypan blue exclusion.</li>
	</ol>
	</li>
	<li>The bone marrow
	<ol>
		<li>Carefully grasp the intact femur using a tissue thumb forceps without fracturing it. Cut off the very end of the proximal femur with a sharp scissor in order to facilitate flushing of the bone. Turn the femur around and position a 23 G needle in the middle of the intercondylar notch of the femur. Whilst applying gentle pressure rotate the needle between thumb and index finger in order to drill a hole in the intercondylar notch to enter the bone cavity. 
		NOTE: Sometimes particles of the bone can obstruct the needle after drilling the hole, to avoid unnecessary high pressure during flushing a change of needle before flushing is recommended.</li>
		<li>Flush the bone with 6 mL of RPMI with 10% FBS (or until the flush turns white) using a 10 mL syringe with a 23 G needle onto a 70 &#181;m cell strainer that is placed on a 15 mL tube. Gently press the bone marrow through the cell strainer with a plunger of a 3 mL syringe and rinse the strainer with another 3 mL of RPMI. 
		NOTE: The bones should appear white once all the marrow has been flushed out completely.</li>
		<li>Spin the cells (500 x g, 5 min, RT) and resuspend the pellet in 5 mL of RBC lysis buffer. Incubate for 5 min at RT and stop the reaction by diluting the lysis buffer with 10 mL of PBS.</li>
		<li>Spin the cells (500 x g, 5 min, RT), discard supernatant and resuspend the pellet in 1 mL of PBS. Refilter the suspension using a 30 &#181;m cell strainer into a new 15 mL tube to remove coagulated cells. Count the number of live cells on a hemocytometer using Trypan blue exclusion.</li>
	</ol>
	</li>
	<li>The synovial tissue
	<ol>
		<li>Dice the two synovial tissue blocks into tiny pieces with a fine surgical scissor. Transfer the samples with medium into a 15 mL tube. Rinse the old well with additional 0.5 mL of RPMI to get remaining cells and synovial tissues, transfer to falcon tube (final volume 2 mL). 
		TIP: Use a transfer pipette and cut of the tip where the diameter widens in this step.</li>
		<li>Reconstitute enzyme and aliquot according to manufacturer instructions (e.g., Liberase). Add sufficient enzyme to result in a final concentration of 1 Unit/mL (a total of 2 Units per sample). Digest at 37 &#176;C for 2 h using a MACS rotator.</li>
		<li>Stop the digestion by adding 8 mL of RPMI with 10% FBS and filter cell suspension through a 70 &#181;m cell strainer into a new 15 mL tube. Rinse the old 15 mL tube with another 5 mL of RPMI with 10% FCS medium and filter cell suspension through same cell strainer into the new tube (15 mL final volume).</li>
		<li>Spin the cells (500 x g, 10 min, RT), discard supernatant and resuspend the pellet in 500 &#181;L of PBS. Count the number of live cells on a hemocytometer using Trypan blue exclusion.</li>
	</ol>
	</li>
</ol>
3. Allocation of cells
<ol>
	<li>Label two 96-well plates (U-bottom shape) with type of tissue, animal ID, and the designated antibody panel. A total of two antibody panels are used in this protocol: Monocyte subset panel (extracellular staining) and T-cell subset panel (extra- and intracellular staining).</li>
	<li>Provide 5 x 105 cells per well using the respective single cell suspensions. 
	NOTE: When setting up the experiment, assess the absolute number of cells that is expected per group and tissue type (treated animals have a higher cell count in tissues than control animals). When resuspending the cell pellet during the last step of generating single cell suspensions, choose an appropriate amount of PBS in order to end up with a concentration of 5 x 105 per 200 &#181;L. The 96-well plate used here can hold a maximum of 300 &#181;L and typically, 200 &#181;L is ideal to minimize the risk of cross-contamination due to spillage.</li>
	<li>For each panel and tissue type, distribute at least 5 x 105 cells as unstained controls in wells that have been clearly marked.</li>
</ol>
4. Monocyte Subset Panel
<ol>
	<li>Perform viability staining: Spin cells (500 x g, 5 min, 4 &#176;C) using a plate spinner and wash the cells once with 200 &#181;L of 1x PBS. Prepare a stock solution of cell-impermeant amine-reactive dye (viability stain) diluted 1:50 in 1x PBS.Thereafter, resuspend the cell pellets with 100 &#181;L of this stock solution resulting in an absolute volume of 2 &#181;L of viability stain per well. Incubate for 15 min at 4 &#176;C protected from light. 
	NOTE: The optimal amount of viability stain needed should be determined by performing a dose titration curve. In addition, diluted viability stain stock solution should be used in a single day and not be stored. Refer to manufacturer&#8217;s instructions for more information on how to reconstitute, dilute and store the viability stain.</li>
	<li>During the incubation, prepare the cocktail of antibodies in an appropriate volume of FACS buffer (Ca2+ and Mg+ free PBS containing 0.1%BSA and 0.02% sodium azide). Wash the cells twice with 200 &#181;L of FACS buffer, centrifuge (500 x g, 5 min, 4 &#176;C) and resuspend each pellet with 100 &#181;L of the antibody mixture or appropriate control mixture. Incubate for 30 min at 4 &#176;C protected from light. 
	NOTE: Please be aware that sodium azide is toxic to cells. In the current protocol the concentration of sodium azide in the flow buffer is very low (0.02%) and samples are run immediately after staining thus, not causing any issue. If downstream functional assays of sorted cells are planned, it might be beneficial to make up fresh FACS buffer each day of experiments and not use any sodium azide. When using a multitude of antibodies, it is advised to add an appropriate amount of &#8220;Brilliant Stain Buffer&#8221; to the cocktail of antibodies to enhance results.</li>
	<li>Wash the cells twice with 200 &#181;L of FACS buffer and resuspend the cells in 250 &#181;L of FACS + EDTA buffer (FACS buffer containing 1 mM EDTA). Transfer samples into labeled FACS tubes. Keep samples at 4 &#176;C and protected from light until acquisition. 
	NOTE: Immune cells have the tendency to be sticky. In order to minimize both the risk of blockage and number of doublets it is recommended to add 1 mM EDTA to the final flow buffer.</li>
</ol>
5. T Cell subset panel
<ol>
	<li>Perform viability staining: Spin cells (500 x g, 5 min, 4 &#176;C) using a plate spinner and wash the cells once with 200 &#181;L of 1x PBS. Prepare a stock solution of cell-impermeant amine-reactive dye (viability stain) diluted 1:50 in 1x PBS. Thereafter, resuspend the cell pellets with 100 &#181;L of this stock solution resulting in an absolute volume of 2 &#181;L of viability stain per well. Incubate for 15 min at 4 &#176;C protected from light.</li>
	<li>Whilst incubating the samples, prepare the extracellular staining antibody cocktail in an appropriate volume of 1x FACS buffer. Wash the cells twice with 200 &#181;L of 1x FACS buffer, spin them down (500 x g, 5 min, 4 &#176;C) and resuspend each pellet with 100 &#181;L of the antibody mixture or appropriate control mixture. Incubate for 30 min at 4 &#176;C protected from light.</li>
	<li>Perform the intracellular staining with a fixation and permeabilization kit following the manufacturer&#8217;s instructions. Wash the cells twice with 200 &#181;L of 1x FACS buffer and resuspend in 200 &#181;L of fixation buffer. Incubate for 40 min at 4 &#176;C protected from light.</li>
	<li>During the incubation, prepare the cocktail of antibodies (intracellular staining) in an appropriate volume of 1x permeabilization and wash buffer. Collect cells by spinning (750 x g, 5 min, 4 &#176;C) and wash cells twice with 200 &#181;L of 1x perm/wash buffer. 
	NOTE: Fixation and permeabilization results in cells that tend to be a bit harder to properly pellet. In order to minimize cell loss during the subsequent washing steps, increase the centrifugal force to 750 x g. Alternatively a longer spinning cycle could also be applied. However, this would result in a considerably longer time that is needed to prepare the cells.</li>
	<li>Spin cells (750 x g, 5 min, 4 &#176;C) and resuspend each pellet with 100 &#181;L of the antibody mixture or appropriate control mixture. Incubate for 40 min at 4 &#176;C protected from light.</li>
	<li>Wash the cells twice with 200 &#181;L of 1x perm/wash buffer and resuspend the cells in 250 &#181;L of FACS + EDTA buffer. Transfer samples into labeled FACS tubes. Keep samples at 4 &#176;C and protected from light until acquisition. 
	NOTE: For each antibody the optimal concentration needs to be determined by performing a dose titration curve. Concentration between antibodies can differ drastically: CD3 and CD80 was used with a dilution factor of 1:1, while CD11b and CD4 was used with a dilution factor of 1:6400. When titrating the antibody concentration use the same number of cells that will be used during the experiments.</li>
</ol>
6. Compensation, appropriate controls and gating
<ol>
	<li>Setting up the experiment
	<ol>
		<li>Once the optimal antibody concentration has been determined run unstained and single stained controls for compensation to adjust for spectral overlap. 
		NOTE: Run all compensation controls with both cells and compensation beads. Use whatever generates the brightest results (highest MFI of positive events) for compensation. MFI stands for mean fluorescence intensity and is often used to describe and define the mean intensity of the generated signal and thus, level of antibody expression.</li>
		<li>Run fluorescence minus one (FMO) controls and isotype controls when starting a new multicolor experiment. Further details regarding FMO have been previously published19.</li>
		<li>Determine the optimal Forward Scatter Area (FSC-A) voltage and Side Scatter Area (SSC-A) voltage in order to detect the leukocyte population in unstained controls of each tissue type. 
		NOTE: The fixation and permeabilization process alters the dimensions of the cell. Thus, the FSC-A and SSC-A voltages for the Monocyte Subset Panel and T Cell Subset Panel differ considerably. In order to find the optimal voltages for the T Cell Subset Panel, use cells that have been single stained with CD3 and back gate towards the leukocyte populations while adjusting the FSC-A and SSC-A values.</li>
	</ol>
	</li>
	<li>Gating strategy
	<ol>
		<li>Once the optimal FSC-A and SSC-A voltage has been determined, set up a primary gate on the leukocyte population. 
		NOTE: Prior to each experiment calibrate the cytometer using calibration beads as per manufacturer&#8217;s instructions and run unstained beads. Leukocyte populations of different time points should have comparable FSC and SSC properties (slight differences between tissue types are expected and normal). If FSC and SSC varies considerable trouble shoot the cytometer and sample generation.</li>
		<li>Exclude doublets: Plot FSC-A (y-axis) and FSC-H (x-axis). Singlets appear as a diagonal of this plot. Gate on singlets.</li>
		<li>Exclude dead cell: Plot FVS510 (viability stain) (x-axis) and FSC-A (y-axis). Dead cells will appear as positive events, thus gate on live cells. 
		NOTE: True negative cells will be visible in unstained controls. Thus, adjust this gate for each set of samples when running the unstained controls prior to stained samples. Further gating depends on the antibody panel and cell type that is investigated. Gating strategies for each panel used in this protocol can be found in Figure 1 and Figure 4, respectively.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+surgical+destabilization+of+the+medial+meniscus+(DMM)+model+of+osteoarthritis+in+the+129/SvEv+mouse.">The surgical destabilization of the medial meniscus (DMM) model of osteoarthritis in the 129/SvEv mouse.</a> Osteoarthritis and Cartilage. 15 (9), 1061-1069 (2007).</li><li>Laroumanie, F., Dale, B. L., Saleh, M. A., Madhur, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+Staining+and+Flow+Cytometry+to+Identify+Lymphocyte+Subsets+within+Murine+Aorta,+Kidney+and+Lymph+Nodes+in+a+Model+of+Hypertension.">Intracellular Staining and Flow Cytometry to Identify Lymphocyte Subsets within Murine Aorta, Kidney and Lymph Nodes in a Model of Hypertension.</a> Journal of Visualized Experiments. (119), e55266 (2017).</li><li>Yu, Y. 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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-invasive+mouse+models+of+post-traumatic+osteoarthritis.">Non-invasive mouse models of post-traumatic osteoarthritis.</a> Osteoarthritis and Cartilage. 23 (10), 1627-1638 (2015).</li><li>Shi, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+and+alteration+of+lymphatic+vessels+in+knee+joints+of+normal+and+osteoarthritic+mice.">Distribution and alteration of lymphatic vessels in knee joints of normal and osteoarthritic mice.</a> Arthritis &amp; Rheumatology. 66 (3), 657-666 (2014).</li><li>Harrell, M. I., Iritani, B. M., Ruddell, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lymph+node+mapping+in+the+mouse.">Lymph node mapping in the mouse.</a> Journal of Immunological Methods. 332 (1-2), 170-174 (2008).</li><li>Zhao, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+the+culture+and+isolation+of+murine+synovial+fibroblasts.">A protocol for the culture and isolation of murine synovial fibroblasts.</a> Biomedical Reports. 5 (2), 171-175 (2016).</li><li>Belluzzi, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contribution+of+Infrapatellar+Fat+Pad+and+Synovial+Membrane+to+Knee+Osteoarthritis+Pain.">Contribution of Infrapatellar Fat Pad and Synovial Membrane to Knee Osteoarthritis Pain.</a> BioMed Research International. 2019, 18 (2019).</li></ol>

The authors have nothing to disclose.

The methods described in this protocol have been designed and tested to reliably identify various subsets from both monocytes/macrophages and T-cells within the murine spleen, bone marrow, lymph nodes, and synovial tissue in a murine model of osteoarthritis (OA). The current protocol can easily be modified to investigate different tissue types, or other cell types by exchanging antibodies, and can be used for alternative murine models of OA. When testing other tissue types, it is critical to test the specificity of each ...

Osteoarthritis (OA) is a debilitating and painful disease involving various pathologies of all tissues associated with the joint1. Affecting approximately 3.8% of the global population2, OA is one of the most prevalent musculoskeletal diseases and it is to become the 4th leading cause of disability worldwide by 20203. Post-traumatic OA occurs after a joint injury and accounts for at least 12% of all OA and up to 25% of OA in susceptible joints such as the knee4,5. Furthermore, joint injury increases the lifetime risk of OA by more than five times6. Not all injuries with apparently similar instability will go on to develop OA, and therefore defining factors that drive the long-term OA-risk remains challenging. It is crucial in order to develop effective treatments to prevent and/or treat post-traumatic OA, to investigate and better define the injury-specific pathology, causes, and mechanisms that predispose to OA1.
OA and its defining cartilage destruction was previously attributed entirely to mechanical stress and, thus, OA was considered a non-inflammatory disease2. However, more recent studies have shown an inflammatory infiltration of synovial membranes and an increase of inflammatory cells in the synovial tissue in patients with OA compared to healthy controls2, shedding light on an inflammatory component as a potential driving force in OA. Further studies indicated that abnormalities in both the CD4+ and CD8+ T-cell profile as well as monocytes/macrophages of the innate immune system may contribute to the pathogenesis of OA2,7. Detailed investigations into these abnormalities revealed relevant roles for various T cell subsets2, such as Th18, Th29, Th178 and T regulatory (Treg) populations10,11. Despite this compelling evidence, the causal relationship between the alteration of T-cell responses and the development and progression of OA is still unknown2.
In addition to specific T-cells having a role in OA, recent studies suggest that differentially polarized/activated macrophages may be associated with pathogenesis of OA12. In particular, macrophages originating from blood monocytes accumulate in the synovium and polarize into either classically activated macrophages (M1) or alternatively activated macrophages (M2) during OA development, implying a correlation between monocyte derived macrophages and OA13. In contrast, certain subsets of macrophages populate organs early during development and self-sustain their numbers in a monocyte independent matter14. Recently, a joint protective function mediated by a tight-junction barrier was shown for these synovial-tissue-resident macrophages (STRMs)14. These findings indicate that abnormalities in particular macrophage subsets may play a crucial role during development of OA. However, the interactions between this inflammatory cellular response and the structural changes in the joint subsequent to trauma is unknown.
Historically, analysis of immune cells in the synovial tissue was restricted to immunohistochemistry (IHC) or mRNA expression by reverse-transcription polymerase chain reaction (RT-PCR) approaches15,16. However, both IHC and RT-PCR lack the ability to identify multiple different cell types and their subsets simultaneously, thus, limiting the applicability of these methods. Furthermore, IHC is limited to analysis of small samples of tissue and may miss focal inflammatory cell accumulations. Over the last several years, a myriad of surface markers for various cell types have been developed, and subsets of immune cells can now be reliably identified by distinct combinations of these markers. Due to steady technical progress, flow cytometers are now capable of identifying a multitude of different fluorochromes simultaneously enabling analysis of large multicolor antibody panels.
Flow cytometry provides investigators with a powerful technique that allows simultaneous identification and quantification of a multitude of immune cells and their subsets at the single cell level. We have developed and optimized both an extracellular staining assay to identify monocytes/macrophages and their subsets as well as an extra/intracellular staining assay to identify T-cells and their subsets within 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 models17.

<table>
	<tbody>
		<tr>
			<td>APC anti-mouse CD194 (CCR4)</td>
			<td>BioLegend</td>
			<td>131212</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Stain Buffer Plus 1000Tst</td>
			<td>BD</td>
			<td>566385</td>
			<td>Buffers</td>
			<td></td>
		</tr>
		<tr>
			<td>Fixable Viability Stain 510, 100 &#181;g</td>
			<td>BD</td>
			<td>564406</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Fixable Viability Stain 510, 100 &#181;g</td>
			<td>BD</td>
			<td>564406</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Liberase, Research Grade</td>
			<td>Roche</td>
			<td>5401127001</td>
			<td>Enzyme for synovial tissue</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD11b APC-R700 M1/70, 100 &#181;g</td>
			<td>BD</td>
			<td>564985</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD11C PE-CF594 HL3, 100 &#181;g</td>
			<td>BD</td>
			<td>562454</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD183 BB700 CXCR3-173, 50 &#181;g</td>
			<td>BD</td>
			<td>742274</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD206 Alexa 647 MR5D3, 25 &#181;g</td>
			<td>BD</td>
			<td>565250</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD25 BV605 PC61, 50 &#181;g</td>
			<td>BD</td>
			<td>563061</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD3e APC-Cy7 145-2C11, 100 &#181;g</td>
			<td>BD</td>
			<td>557596</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD4 PE-Cy7 RM4-5, 100 &#181;g</td>
			<td>BD</td>
			<td>552775</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD44 APC-R700 IM7, 50 &#181;g</td>
			<td>BD</td>
			<td>565480</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD62L BB515 MEL-14, 100 &#181;g</td>
			<td>BD</td>
			<td>565261</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD69 BV711 H1.2F3, 50 &#181;g</td>
			<td>BD</td>
			<td>740664</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD80 BV650 16-10A1, 50 &#181;g</td>
			<td>BD</td>
			<td>563687</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms CD8a BV786 53-6.7, 50 &#181;g</td>
			<td>BD</td>
			<td>563332</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms F4/80 BV421 T45-2342, 50 &#181;g</td>
			<td>BD</td>
			<td>565411</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms Foxp3 PE MF23, 100 &#181;g</td>
			<td>BD</td>
			<td>560408</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms I-A I-E BV711 M5/114.15.2, 50 &#181;g</td>
			<td>BD</td>
			<td>563414</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms Ly-6C PE-Cy7 AL-21, 50 &#181;g</td>
			<td>BD</td>
			<td>560593</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms Ly-6G APC-Cy7 1A8, 50 &#181;g</td>
			<td>BD</td>
			<td>560600</td>
			<td>Monocyte Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms NK1.1 BV650 PK136, 50 &#181;g</td>
			<td>BD</td>
			<td>564143</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Ms ROR Gamma T BV421 Q31-378, 50 &#181;g</td>
			<td>BD</td>
			<td>562894</td>
			<td>T-Cell Panel</td>
			<td></td>
		</tr>
		<tr>
			<td>Red Blood Cell Lysing Buffer</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Buffers</td>
			<td>Description in: Immune Cell Isolation from Mouse Femur Bone Marrow / Xiaoyu Liu and Ning Quan/ Bio Protoc. 2015 October 20; 5(20): .</td>
		</tr>
		<tr>
			<td>Transcription Factor Buffer Set 100Tst</td>
			<td>BD</td>
			<td>562574</td>
			<td>Buffers</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Representative results from both the monocyte subset panel and T-cell subset panel are described below.
Figure 1 illustrates the hierarchical gating strategy for the monocyte subset panel on immune cells gathered from bone marrow of DMM treated animals. The same strategy was used and verified in all other tissue types. When setting up the experiment, the Forward Scatter Area (FSC-A) and Side Scatter Area (SSC-A) voltage was determined for each tissue type to ident...

Watch this Scientific Journal Video about Flow Cytometry Analysis of Immune Cell Subsets within the Murine Spleen, Bone Marrow, Lymph Nodes and Synovial Tissue in an Osteoarthritis Model at JoVE.com

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

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.

Osteoarthritis (OA) is one of the most prevalent musculoskeletal diseases, affecting patients suffering from pain and physical limitations. Recent ...

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19.2K 次观看.  University of Sydney. 我们的协议能够详细识别和分析单细胞和巨噬细胞和T细胞及其相关的子集内的神经元和卵巢和各种其他组织的免疫系统。我们的技术有助于从合成器和其他相关组织中可靠地收获生命免疫细胞，以便全面描述骨关节炎免疫反应。对于从骨关节炎模型小鼠分离感兴趣的组织，用鼠标在超活位置，用70%乙醇擦拭胸部、腹部和腿部。使用剪刀小心地打开腹部中线的皮肤，同时...

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

Using Quantitative Real-time PCR to Determine Donor Cell Engraftment in a Competitive Murine Bone Marrow Transplantation Model

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules that regulate HSCs. In these transplant model systems, the function of HSCs is determined by the ability of these cells to engraft and reconstitute lethally irradiated recipient mice. Commonly, the donor cell contribution/engraftment is measured by antibodies to donor- specific cell surface proteins using flow cytometry. However, this method heavily depends on the specificity and the ability of the cell surface marker to differentiate donor-derived cells from recipient-originated cells, which may not be available for all mouse strains. Considering the various backgrounds of genetically modified mouse strains in the market, this cell surface/ flow cytometry-based method has significant limitations especially in mouse strains that lack well-defined surface markers to separate donor cells from congenic recipient cells. Here, we reported a PCR-based technique to determine donor cell engraftment/contribution in transplant recipient mice. We transplanted male donor bone marrow HSCs to lethally irradiated congenic female mice. Peripheral blood samples were collected at different time points post transplantation. Bone marrow samples were obtained at the end of the experiments. Genomic DNA was isolated and the Y chromosome specific gene, Zfy1, was amplified using quantitative Real time PCR. The engraftment of male donor-derived cells in the female recipient mice was calculated against standard curve with known percentage of male vs. female DNAs. Bcl2 was used as a reference gene to normalize the total DNA amount. Our data suggested that this approach reliably determines donor cell engraftment and provides a useful, yet simple method in measuring hematopoietic cell reconstitution in murine bone marrow transplantation models. Our method can be routinely performed in most laboratories because no costly equipment such as flow cytometry is required.

Determining donor cell engraftment presents a challenge in mouse bone marrow transplant models that lack well-defined phenotypical markers. We described a methodology to quantify male donor cell engraftment in female transplant recipient mice. This method can be used in all mouse strains for the study of HSC functions.

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules ...

High-throughput Flow Cytometry Cell-based Assay to Detect Antibodies to N-Methyl-D-aspartate Receptor or Dopamine-2 Receptor in Human Serum

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases in children and adults. These antibodies have been used to guide diagnosis and treatment. Cell-based assays have improved the detection of antibodies in patient serum. They are based on the surface expression of brain antigens on eukaryotic cells, which are then incubated with diluted patient sera followed by fluorochrome-conjugated secondary antibodies. After washing, secondary antibody binding is then analyzed by flow cytometry. Our group has developed a high-throughput flow cytometry live cell-based assay to reliably detect antibodies against specific neurotransmitter receptors. This flow cytometry method is straight forward, quantitative, efficient, and the use of a high-throughput sampler system allows for large patient cohorts to be easily assayed in a short space of time. Additionally, this cell-based assay can be easily adapted to detect antibodies to many different antigenic targets, both from the central nervous system and periphery. Discovering additional novel antibody biomarkers will enable prompt and accurate diagnosis and improve treatment of immune-mediated disorders.

Over the recent years, live cell-based assays have been used successfully to detect antibodies against surface and conformational antigens. Here, we describe a method using high-throughput flow cytometry enabling the analysis of large cohorts of patients. Detection of novel antibodies will improve diagnosis and treatment of immune-mediated disorders.

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases ...

A Three-dimensional Tissue Culture Model to Study Primary Human Bone Marrow and its Malignancies

Tissue culture has been an invaluable tool to study many aspects of cell function, from normal development to disease. Conventional cell culture methods rely on the ability of cells either to attach to a solid substratum of a tissue culture dish or to grow in suspension in liquid medium. Multiple immortal cell lines have been created and grown using such approaches, however, these methods frequently fail when primary cells need to be grown ex vivo. Such failure has been attributed to the absence of the appropriate extracellular matrix components of the tissue microenvironment from the standard systems where tissue culture plastic is used as a surface for cell growth. Extracellular matrix is an integral component of the tissue microenvironment and its presence is crucial for the maintenance of physiological functions such as cell polarization, survival, and proliferation. Here we present a 3-dimensional tissue culture method where primary bone marrow cells are grown in extracellular matrix formulated to recapitulate the microenvironment of the human bone (rBM system). Embedded in the extracellular matrix, cells are supplied with nutrients through the medium supplemented with human plasma, thus providing a comprehensive system where cell survival and proliferation can be sustained for up to 30 days while maintaining the cellular composition of the primary tissue. Using the rBM system we have successfully grown primary bone marrow cells from normal donors and patients with amyloidosis, and various hematological malignancies. The rBM system allows for direct, in-matrix real time visualization of the cell behavior and evaluation of preclinical efficacy of novel therapeutics. Moreover, cells can be isolated from the rBM and subsequently used for in vivo transplantation, cell sorting, flow cytometry, and nucleic acid and protein analysis. Taken together, the rBM method provides a reliable system for the growth of primary bone marrow cells under physiological conditions.

In standard culture methods cells are taken out of their physiological environment and grown on the plastic surface of a dish. To study the behavior of primary human bone marrow cells we created a 3-D culture system where cells are grown under conditions recapitulating the native microenvironment of the tissue.

Tissue culture has been an invaluable tool to study many aspects of cell function, from normal development to disease. Conventional cell culture methods ...

High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal systems. HFUS has the ability to detect structures as small as 30 &#181;m, a property that has been utilized for visualizing superficial lymph nodes in rodents in brightness (B)-mode. Combining power Doppler with B-mode imaging allows for measuring circulatory blood flow within lymph nodes and other organs. While HFUS has been utilized for lymph node imaging in a number of mouse&#160; model systems, a detailed protocol describing HFUS imaging and characterization of the cervical lymph nodes in mice has not been reported. Here, we show that HFUS can be adapted to detect and characterize cervical lymph nodes in mice. Combined B-mode and power Doppler imaging can be used to detect increases in blood flow in immunologically-enlarged cervical nodes. We also describe the use of B-mode imaging to conduct fine needle biopsies of cervical lymph nodes to retrieve lymph tissue for histological&#160; analysis. Finally, software-aided steps are described to calculate changes in lymph node volume and to visualize changes in lymph node morphology following image reconstruction. The ability to visually monitor changes in cervical lymph node biology over time provides a simple and powerful technique for the non-invasive monitoring of cervical lymph node alterations in preclinical mouse models of oral cavity disease.

This protocol describes the application of high-frequency ultrasound (HFUS) for imaging mouse cervical lymph nodes. This technique optimizes visualization and quantification of cervical lymph node morphology, volume and blood flow. Image-guided biopsy of cervical lymph nodes and processing of lymph tissue for histological evaluation is also demonstrated.

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal ...

Imaging- and Flow Cytometry-based Analysis of Cell Position and the Cell Cycle in 3D Melanoma Spheroids

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to traditional two-dimensional (2D) cell culture. The spheroid model is able to mimic the effects of cell-cell interaction, hypoxia and nutrient deprivation, and drug penetration. One characteristic of this model is the development of a necrotic core, surrounded by a ring of G1 arrested cells, with proliferating cells on the outer layers of the spheroid. Of interest in the cancer field is how different regions of the spheroid respond to drug therapies as well as genetic or environmental manipulation. We describe here the use of the fluorescence ubiquitination cell cycle indicator (FUCCI) system along with cytometry and image analysis using commercial software to characterize the cell cycle status of cells with respect to their position inside melanoma spheroids. These methods may be used to track changes in cell cycle status, gene/protein expression or cell viability in different sub-regions of tumor spheroids over time and under different conditions.

We describe two complementary methods using the fluorescence ubiquitination cell cycle indicator (FUCCI) and image analysis or flow cytometry to identify and isolate cells in the inner G1 arrested and outer proliferating regions of 3D spheroids.

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to ...

Isolation and Cellular Phenotyping of Mesenchymal Stem Cells Derived from Synovial Fluid and Bone Marrow of Minipigs

Mesenchymal stem cells (MSCs) have been established after isolation from various tissue sources, including bone marrow and synovial fluid. Recently, synovial-fluid-derived MSCs were reported to have multi-lineage differentiation potential and immunomodulatory features, which indicates that these cells can be used for tissue engineering and systemic treatments. This study presents a protocol for simple and non-invasive isolation of MSCs derived from the bone marrow and synovial fluid of minipigs to analyze cell surface markers for cell phenotyping and in vitro culturing. Using sexually mature six-month-old minipigs, bone marrow was extracted from the iliac crest bone using a bone marrow extractor, and the synovial fluid was aspirated from the femorotibial joint. Procedures for the collection of samples from both sources were non-invasive. The protocols for effective isolation of MSCs from harvested cell sources and for creating in vitro culture conditions to expand stable MSCs from minipigs and the application of systemic autologous treatments are provided. For cell phenotyping, the cell surface markers of both cells were analyzed using flow cytometry. In the results, the MSCs were isolated from the synovial fluid of the minipigs and showed that synovial-fluid-derived MSCs have a similar morphology and cell phenotype to bone-marrow-derived MSCs. Therefore, non-invasively obtained synovial fluid is a valuable source of MSCs.

A protocol establishing mesenchymal stem cells (MSCs) isolated from the synovial fluid and bone marrow of minipigs and non-invasively collected using syringe aspiration is presented. Cellular phenotyping was performed using flow cytometry after cell isolation and in vitro cultivation of bone marrow and synovial fluids derived from MSCs.

Mesenchymal stem cells (MSCs) have been established after isolation from various tissue sources, including bone marrow and synovial fluid. Recently, ...

Laparoscopic Technique for Serial Collection of Liver and Mesenteric Lymph Nodes in Macaques

The mesenteric lymph nodes (MLN) and the liver are exposed to microbes and microbial products from the gastrointestinal (GI) tract, making them immunologically unique. The GI tract and associated MLN are sites of early viral replication in human immunodeficiency virus (HIV) infection and the MLN are likely important reservoir sites that harbor latently-infected cells even after prolonged antiretroviral therapy (ART). The liver has been shown to play a significant role in immune responses to lentiviruses and appears to play a significant role in clearance of virus from circulation. Nonhuman primate (NHP) models for HIV and Acquired Immunodeficiency Syndrome (AIDS) closely mimic these aspects of HIV infection and serial longitudinal sampling of primary sites of viral replication and the associated immune responses in this model will help to elucidate critical events in infection, pathogenesis, and the impact of various intervention strategies on these events. Current published techniques to sample liver and MLN together involve major surgery and/or necropsy, which limits the ability to investigate these important sites in a serial fashion in the same animal. We have previously described a laparoscopic technique for collection of MLN. Here, we describe a minimally invasive laparoscopic technique for serial longitudinal sampling of liver and MLN through the same two port locations required for the collection of MLN. The use of the same two ports minimizes the impact to the animals as no additional incisions are required. This technique can be used with increased sampling frequency compared to major abdominal surgery and reduces the potential for surgical complications and associated local and systemic inflammatory responses that could complicate interpretation of results. This procedure has potential to facilitate studies involving NHP models while improving animal welfare.

Here, we describe a minimally invasive laparoscopic technique for serial sampling of liver and mesenteric lymph nodes (MLN) in macaques that allows for increased sampling frequency, and reduces the potential for surgical complications when compared to performing a laparotomy.

The mesenteric lymph nodes (MLN) and the liver are exposed to microbes and microbial products from the gastrointestinal (GI) tract, making them ...

Characterization of Immune Cells in Human Adipose Tissue by Using Flow Cytometry

Infiltration of immune cells in the subcutaneous and visceral adipose tissue (AT) deposits leads to a low-grade inflammation contributing to the development of obesity-associated complications such as type 2 diabetes. To quantitatively and qualitatively investigate the immune cell subsets in human AT deposits, we have developed a flow cytometry approach. The stromal vascular fraction (SVF), containing the immune cells, is isolated from subcutaneous and visceral AT biopsies by collagenase digestion. Adipocytes are removed after centrifugation. The SVF cells are stained for multiple membrane-bound markers selected to differentiate between immune cell subsets and analyzed using flow cytometry. As a result of this approach, pro- and anti-inflammatory macrophage subsets, dendritic cells (DCs), B-cells, CD4+ and CD8+ T-cells, and NK cells can be detected and quantified. This method gives detailed information about immune cells in AT and the amount of each specific subset. Since there are numerous fluorescent antibodies available, our flow cytometry approach can be adjusted to measure various other cellular and intracellular markers of interest.

This article describes a method to analyze immune cell content of adipose tissue by isolation of immune cells from adipose tissue and subsequent analysis using flow cytometry.

Infiltration of immune cells in the subcutaneous and visceral adipose tissue (AT) deposits leads to a low-grade inflammation contributing to the ...

Database-guided Flow-cytometry for Evaluation of Bone Marrow Myeloid Cell Maturation

A working group initiated within the French Cytometry Association (AFC) was developed in order to harmonize the application of multiparameter flow cytometry (MFC) for myeloid disease diagnosis in France. The protocol presented here was agreed-upon and applied between September 2013 and November 2015 in six French diagnostic laboratories (University Hospitals of Saint-Etienne, Grenoble, Clermont-Ferrand, Nice, and Lille and Institut Paoli-Calmettes in Marseille) and allowed the standardization of bone marrow sample preparation and data acquisition. Three maturation databases were developed for neutrophil, monocytic, and erythroid lineages with bone marrow from "healthy" donor individuals (individuals without any evidence of a hematopoietic disease). A robust method of analysis for each myeloid lineage should be applicable for routine diagnostic use. New cases can be analyzed in the same manner and compared against the usual databases. Thus, quantitative and qualitative phenotypic abnormalities can be identified and those above 2SD compared with data of normal bone marrow samples should be considered indicative of pathology. The major limitation is the higher variability between the data achieved using the monoclonal antibodies obtained with the methods based on hybridoma technologies and currently used in clinical diagnosis. Setting criteria for technical validation of the data acquired may help improve the utility of MFC for MDS diagnostics. The establishment of these criteria requires analysis against a database. The reduction of investigator subjectivity in data analysis is an important advantage of this method.

The MDS diagnosis is difficult in the absence of morphological criteria or non-informative cytogenetics. MFC could help refine the MDS diagnostic process. To become useful for clinical practice, the MFC analysis must be based on parameters with sufficient specificity and sensitivity, and data should be reproducible between different operators.

A working group initiated within the French Cytometry Association (AFC) was developed in order to harmonize the application of multiparameter flow ...

Quality-Controlled Sputum Analysis by Flow Cytometry

Sputum, widely used to study the cellular content and other microenvironmental features to understand the health of the lung, is traditionally analyzed using cytology-based methodologies. Its utility is limited because reading the slides is time-consuming and requires highly specialized personnel. Moreover, extensive debris and the presence of too many squamous epithelial cells (SECs), or cheek cells, often renders a sample inadequate for diagnosis. In contrast, flow cytometry allows for high-throughput phenotyping of cellular populations while simultaneously excluding debris and SECs.
The protocol presented here describes an efficient method to dissociate sputum into a single cell suspension, antibody stain and fix cellular populations, and acquire samples on a flow cytometric platform. A gating strategy that describes the exclusion of debris, dead cells (including SECs) and cell doublets is presented here. Further, this work also explains how to analyze viable, single sputum cells based on a cluster of differentiation (CD)45 positive and negative populations to characterize hematopoietic and epithelial lineage subsets. A quality control measure is also provided by identifying lung-specific macrophages as evidence that a sample is derived from the lung and is not saliva. Finally, it has been demonstrated that this method can be applied to different cytometric platforms by providing sputum profiles from the same patient analyzed on three flow cytometers; Navios EX, LSR II, and Lyric. Furthermore, this protocol can be modified to include additional cellular markers of interest. A method to analyze an entire sputum sample on a flow cytometric platform is presented here that makes sputum amenable for developing high-throughput diagnostics of lung disease.

This protocol describes an efficient method for dissociating sputum into a single cell suspension and the subsequent characterization of cellular subsets on standard flow cytometric platforms.

Sputum, widely used to study the cellular content and other microenvironmental features to understand the health of the lung, is traditionally analyzed ...