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

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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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The Lymphatic and Immune System

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19.4K Views.  University of Sydney. Ons protocol maakt de gedetailleerde identificatie en analyse van monocyten en macrofagen en T-cellen en hun relevante subsets binnen de neuronen en ovium en diverse andere weefsels van het immuunsysteem mogelijk. Onze techniek vergemakkelijkt de betrouwbare oogst van leven immuuncellen uit het synovium en andere relevante weefsels voor een uitgebreide karakterisering van de osteoarthritische immuunrespons. Voor de isolatie van weefsels van belang van een artrose model muis, met de muis in ee...

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

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 ...

Immunology and Infection

Intracellular Staining and Flow Cytometry to Identify Lymphocyte Subsets within Murine Aorta, Kidney and Lymph Nodes in a Model of Hypertension

It is now well known that T lymphocytes play a critical role in the development of several cardiovascular diseases1,2,3,4,5. For example, studies from our group have shown that hypertension is associated with an excessive accumulation of T cells in the vessels and kidney during the development of experimental hypertension6. Once in these tissues, T cells produce several cytokines that affect both vascular and renal function leading to vasoconstriction and sodium and water retention1,2. To fully understand how T cells cause cardiovascular and renal diseases, it is important to be able to identify and quantify the specific T cell subsets present in these tissues. T cell subsets are defined by a combination of surface markers, the cytokines they secrete, and the transcription factors they express. The complexity of the T cell population makes flow cytometry and intracellular staining an invaluable technique to dissect the phenotypes of the lymphocytes present in tissues. Here, we provide a detailed protocol to identify the surface and intracellular markers (cytokines and transcription factors) in T cells isolated from murine kidney, aorta and aortic draining lymph nodes in a model of angiotensin II induced hypertension. The following steps are described in detail: isolation of the tissues, generation of the single cell suspensions, ex vivo stimulation, fixation, permeabilization and staining. In addition, several fundamental principles of flow cytometric analyses including choosing the proper controls and appropriate gating strategies are discussed.

This article provides detailed methodology to identify and quantify functional T lymphocyte subsets present within murine kidney, aorta and lymph nodes by intracellular staining and flow cytometry. The model of angiotensin II induced hypertension was chosen to explain, step-by-step, the procedures and fundamental principles of flow cytometry and intracellular staining.

It is now well known that T lymphocytes play a critical role in the development of several cardiovascular diseases1,2,3,4,5. For example, studies from our ...

Creation of a Knee Joint-on-a-Chip for Modeling Joint Diseases and Testing Drugs

The high prevalence of debilitating joint diseases like osteoarthritis (OA) poses a high socioeconomic burden. Currently, the available drugs that target joint disorders are mostly palliative. The unmet need for effective disease-modifying OA drugs (DMOADs) has been primarily caused by the absence of appropriate models for studying the disease mechanisms and testing potential DMOADs. Herein, we describe the establishment of a miniature synovial joint-mimicking microphysiological system (miniJoint) comprising adipose, fibrous, and osteochondral tissue components derived from human mesenchymal stem cells (MSCs). To obtain the three-dimensional (3D) microtissues, MSCs were encapsulated in photocrosslinkable methacrylated gelatin before or following differentiation. The cell-laden tissue constructs were then integrated into a 3D-printed bioreactor, forming the miniJoint. Separate flows of osteogenic, fibrogenic, and adipogenic media were introduced to maintain the respective tissue phenotypes. A commonly shared stream was perfused through the cartilage, synovial, and adipose tissues to enable tissue crosstalk. This flow pattern allows the induction of perturbations in one or more of the tissue components for mechanistic studies. Furthermore, potential DMOADs can be tested via either &#34;systemic administration&#34; through all the medium streams or &#34;intraarticular administration&#34; by adding the drugs to only the shared &#34;synovial fluid&#34;-simulating flow. Thus, the miniJoint can serve as a versatile in vitro platform for efficiently studying disease mechanisms and testing drugs in personalized medicine.

We provide detailed methods for generating four types of tissues from human mesenchymal stem cells, which are used to recapitulate the cartilage, bone, fat pad, and synovium in the human knee joint. These four tissues are integrated into a customized bioreactor and connected through microfluidics, thus generating a knee joint-on-a-chip.

The high prevalence of debilitating joint diseases like osteoarthritis (OA) poses a high socioeconomic burden. Currently, the available drugs that target ...

Bioengineering

Differentiation of Functional Osteoclasts from Human Peripheral Blood CD14+ Monocytes

Osteoclasts (OCs) are bone-resorbing cells that play a pivotal role in skeletal development and adult bone remodeling. Several bone disorders are caused by increased differentiation and activation of OCs, so the inhibition of this pathobiology is a key therapeutic principle.Two key factors drive the differentiation of OCs from myeloid precursors: macrophage colony-stimulating factor (M-CSF) and receptor activator of nuclear factor kappa-B ligand (RANKL). Human circulating CD14+ monocytes have long been known to differentiate into OCs in vitro. However, the exposure time and the concentration of RANKL influence the differentiation efficiency. Indeed, protocols for the generation of human OCs in vitro have been described, but they often result in a poor and lengthy differentiation process. Herein, a robust and standardized protocol for generating functionally active mature human OCs in a timely manner is provided. CD14+ monocytes are enriched from human peripheral blood mononuclear cells (PBMCs) and primed with M-CSF to upregulate RANK. Subsequent exposure to RANKL generates OCs in a dose- and time-dependent manner. OCs are identified and quantified by staining with tartrate acid-resistant phosphatase (TRAP) and light microscopy analysis. Immunofluorescence staining of nuclei and F-actin is used to identify functionally active OCs. In addition, OSCAR+CD14&#8722; mature OCs are further enriched via flow cytometry cell sorting, and OC functionality quantified by mineral (or dentine/bone) resorption assays and actin ring formation. Finally, a known OC inhibitor, rotenone, is used on mature OCs, demonstrating that adenosine triphosphate (ATP) production is essential for actin ring integrity and OC function. In conclusion, a robust assay for differentiating high numbers of OCs is established in this work, which in combination with actin ring staining and an ATP assay&#160;provides a useful in vitro model to evaluate OC function and to screen for novel therapeutic compounds that can modulate the differentiation process.

Differentiation of Functional Osteoclasts from Human Peripheral Blood CD14+ Monocytes

Osteoclasts are key bone-resorbing cells in the body. This protocol describes a reliable method for the in vitro differentiation of osteoclasts from human peripheral blood monocytes. This method can be used as an important tool to further understand osteoclast biology in homeostasis and in diseases.

Osteoclasts (OCs) are bone-resorbing cells that play a pivotal role in skeletal development and adult bone remodeling. Several bone disorders are caused ...

Standardized Histomorphometric Evaluation of Osteoarthritis in a Surgical Mouse Model

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, primarily in the hip and knee joints, which results in significant impacts on patient mobility and quality of life. To date, there are no existing curative therapies for OA able to slow down or inhibit cartilage degeneration. Presently, there is an extensive body of ongoing research to understand OA pathology and discover novel therapeutic approaches or agents that can efficiently slow down, stop, or even reverse OA. Thus, it is crucial to have a quantitative and reproducible approach to accurately evaluate OA-associated pathological changes in the joint cartilage, synovium, and subchondral bone. Currently, OA severity and progression are primarily assessed using the Osteoarthritis Research Society International (OARSI) or Mankin scoring systems. In spite of the importance of these scoring systems, they are semiquantitative and can be influenced by user subjectivity. More importantly, they fail to accurately evaluate subtle, yet important, changes in the cartilage during the early disease states or early treatment phases. The protocol we describe here uses a computerized and semiautomated histomorphometric software system to establish a standardized, rigorous, and reproducible quantitative methodology for the evaluation of joint changes in OA. This protocol presents a powerful addition to the existing systems and allows for more efficient detection of pathological changes in the joint.

The current protocol establishes a rigorous and reproducible method for quantification of morphological joint changes that accompany osteoarthritis. Application of this protocol can be valuable in monitoring disease progression and evaluating therapeutic interventions in osteoarthritis.

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, ...

Destabilization of the Medial Meniscus and Cartilage Scratch Murine Model of Accelerated Osteoarthritis

Osteoarthritis is the most prevalent musculoskeletal disease in people over 45, leading to an increasing economic and societal cost. Animal models are used to mimic many aspects of the disease. The present protocol describes the destabilization and cartilage scratch model (DCS) of post-traumatic osteoarthritis. Based on the widely used destabilization of the medial meniscus (DMM) model, DCS introduces three scratches on the cartilage surface. The current article highlights the steps to destabilize the knee by transecting the medial meniscotibial ligament followed by three intentional superficial scratches on the articular cartilage. The possible analysis methods by dynamic weight-bearing, microcomputed tomography, and histology are also demonstrated. While the DCS model is not recommended for studies that focus on the effect of osteoarthritis on the cartilage, it enables the study of osteoarthritis development in a shorter time window, with special focus on (1) osteophyte formation, (2) osteoarthritic and injury pain, and (3) the effect of cartilage damage in the whole joint.

The present protocol describes the controlled microblade scratches on the surface of the articular cartilage after destabilizing the mouse knee by cutting the medial miniscotibial ligament. This animal model presents an accelerated form of osteoarthritis (OA) suitable for studying osteophyte formation, osteosclerosis, and early-stage pain.

Osteoarthritis is the most prevalent musculoskeletal disease in people over 45, leading to an increasing economic and societal cost. Animal models are ...

Synovial Fluid Analysis to Identify Osteoarthritis

Synovial fluid (SF) analysis is important in diagnosing osteoarthritis (OA). Macroscopic and microscopic features, including total and differential white blood cell (WBC) count, help define the non-inflammatory nature of SF, which is a hallmark of OA. In patients with OA, WBC in SF samples usually does not exceed 2000 cells per microliter, and the percentage of inflammatory cells, such as neutrophils, is very low or absent. Calcium crystals are frequent in SF collected from OA patients. Although their role in the pathogenesis of OA remains unclear, they have been associated with a mild inflammatory process and a more severe disease progression. Recently, calcium crystals have been described in both the early and late stages of OA, indicating that they may play a vital role in diagnosing different clinical subsets of OA and pharmacological treatment. The overall goal of SF analysis in OA is two-fold: to ascertain the non-inflammatory degree of SF and to highlight the presence of calcium crystals.

Synovial fluid analysis under transmitted, polarized, and compensated light microscopy is used to evaluate the inflammatory or non-inflammatory nature of a sample through simple steps. It is particularly useful in osteoarthritis to detect calcium crystals and identify a more severe subset of osteoarthritis.

Synovial fluid (SF) analysis is important in diagnosing osteoarthritis (OA). Macroscopic and microscopic features, including total and differential white ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...

Biology

Ameliorating Osteoarthritis in Mice Using Silver Nanoparticles

Knee osteoarthritis (KOA) is one of the most commonly encountered degenerative diseases of the joints in people over 45 years of age. Currently, there are not any effective therapeutics for KOA,and the only end-point strategy is total knee arthroplasty (TKA); therefore, KOA is associated with economic burdens and societal costs. The immune inflammatory response is involved in the occurrence and development of KOA. We previously established a mouse model of KOA using type II collagen. Hyperplasia of the synovial tissue was present in the model, alongside a large number of infiltrated inflammatory cells. Silver nanoparticles have substantial anti-inflammatory effects and have been widely used in tumor therapy and surgical drug delivery. Therefore, we evaluated the therapeutic effects of silver nanoparticles in a collagenase II-induced KOA model. The experimental results showed that silver nanoparticles significantly reduced synovial hyperplasia and the infiltration of neutrophils in the synovial tissue. Hence, this work demonstrates the identification of a novel strategy for OA and provides a theoretical basis for preventing the progress of KOA.

Presented here is a protocol for using silver nanoparticles to effectively ameliorate the acute symptoms of type II collagenase-induced osteoarthritis mice, including synovial inflammation, synovial hyperplasia, vascular hyperplasia, etc.

Knee osteoarthritis (KOA) is one of the most commonly encountered degenerative diseases of the joints in people over 45 years of age. Currently, there are ...

Unveiling Inflammatory Arthritis&#58; Exploring Functions of Synovial Cells

Isolation and Culture of Primary Synovial Macrophages and Fibroblasts from Murine Arthritis Tissue

Rheumatoid arthritis is an autoimmune disease that leads to chronic inflammation of joints. Synovial macrophages and synovial fibroblasts have central roles in the pathogenesis of rheumatoid arthritis. It is important to understand the functions of both cell populations to reveal the mechanisms underlying pathological progression and remission in inflammatory arthritis. In general, in vitro experimental conditions should mimic the in vivo environment as much as possible. Primary tissue-derived cells have been used in experiments characterizing synovial fibroblasts in arthritis. In contrast, in experiments investigating the biological functions of macrophages in inflammatory arthritis, cell lines, bone marrow-derived macrophages, and blood monocyte-derived macrophages have been used. However, it is unclear whether such macrophages actually reflect the functions of tissue-resident macrophages. To obtain resident macrophages, previous protocols were modified to isolate and expand both primary macrophages and fibroblasts from synovial tissue in an inflammatory arthritis mouse model. These primary synovial cells may be useful for in vitro analysis of inflammatory arthritis.

Author Spotlight: Isolation and Culture of Primary Synovial Macrophages and Fibroblasts from Murine Arthritis Tissue  

The present study provides a modified protocol to isolate synovial macrophages and fibroblasts from murine inflammatory arthritis tissue.

Rheumatoid arthritis is an autoimmune disease that leads to chronic inflammation of joints. Synovial macrophages and synovial fibroblasts have central ...

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

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