This work is supported by NIH grants AI125701 and AI139721, Cancer Research Institute CLIP program and American Cancer Society grant RSG-18-222-01-LIB to N.Z. We thank Karla Gorena and Sebastian Montagnino from Flow Cytometry Facility. Data generated in the Flow Cytometry Shared Resource Facility were supported by the University of Texas Health Science Center at San Antonio (UTHSCSA), NIH/NCI grant P30 CA054174-20 (Clinical and Translational Research Center [CTRC] at UTHSCSA), and UL1 TR001120 (Clinical and Translational Science Award).

All animal experiments performed following this protocol must be approved by the respective institutional Animal Care and Use Committee (IACUC). All procedures described here have been approved by IACUC UT Health San Antonio.
1. Adoptive transfer of P14 T cells into C57BL/6 recipients and LCMV infection
<ol>
	<li>Use P14 mice at 6-12 weeks of age. Ensure that the sex of donor P14 TCR transgenic mouse should match the sex of C57BL/6 recipient mice. Otherwise, cells such as male T cells transferring into female mice will result in immune-mediated rejection of circulating donor T cells in around 12 days14.</li>
	<li>Purify na&#239;ve CD8+ T cells from the spleen and lymph nodes of a P14 TCR transgenic mouse using mouse CD8+ T cell purification kit following the manufacturer&#8217;s protocol.
	<ol>
		<li>Briefly, dissect spleen and lymph nodes, mash the samples to generate single cell suspension.</li>
		<li>To enrich for na&#239;ve T cells, add biotin-anti-CD44 antibody during the first antibody incubation step13. 
		NOTE: Negative selection commercial kit used in this step contains two major components for two steps of incubation (i.e., biotin-antibody mixture incubation and streptavidin-magnetic bead incubation).</li>
	</ol>
	</li>
	<li>Use a hemocytometer to count the cells and inject 1,000 to 10,000 purified P14 T cells in 200 &#181;L of PBS into each sex matched C57BL/6 recipient via the tail vein. 
	NOTE: Detailed tail vein injection protocol is described in step 2.</li>
	<li>On the next day, all C57BL/6 recipients receive 2 x 105 pfu LCMV Armstrong diluted in 200 &#181;L of PBS via an intraperitoneal route.</li>
</ol>
2. Intravascular labeling of CD8+ T cells
NOTE: Depending on the research interests, different time points can be chosen following infection. An example of day 30 post infection (day 30 after step 1.4) is demonstrated, which is often considered as an early memory time point for CD8 T cell response.
<ol>
	<li>Dilute biotin anti-CD8&#945; antibody to 15 &#181;g/mL in PBS. Ensure each mouse receive 200 &#956;L of PBS containing 3 &#181;g antibody. 
	NOTE: Biotin anti-CD8 antibody is used here so that it will be more flexible to design the final FACS staining panel. Fluorescent dye-labeled antibody can be used directly. However, it is important to use different clones of monoclonal antibodies for intravenous labeling versus FACS staining.</li>
	<li>Properly heat the tail vein of mice with an overhead heat lamp for 5-10 min to dilate the veins. 
	NOTE: Extra care must be taken to prevent overheating the animals. Reduce the heat or remove the heat lamp if the mice stopped to move around.</li>
	<li>Draw 200 &#181;L of prediluted anti-CD8&#945; antibody mix into a 100 U insulin syringe (28G) and remove any air bubbles by moving the piston up and down. Bend the needle to create a 150&#176; angle between the needle and syringe, bevel up, so the needle will be parallel to the vein.</li>
	<li>Restrain the mouse with a rodent restrainer of appropriate size and spray the tail with 70% ethanol to make the vein clearly visible. 
	NOTE: The duration of the restraint should be kept to a minimum.</li>
	<li>Hold the tail at the distal end with thumb and middle fingers of the non-dominant hand. Place the index finger underneath the site where the needle will be inserted.</li>
	<li>Hold the syringe with the dominant hand and insert the needle into the vein in parallel towards the direction of the heart. 
	NOTE: When placing correctly, the needle should move smoothly into the vein.</li>
	<li>Slowly inject 200 &#181;L of anti-CD8&#945; antibody via the tail vein. 
	NOTE: If there is a resistance or a white area is seen above the needle on the tail, the needle is not inside the vein. Start the initial injection close to the tip of the tail. When necessary, move forward towards the bottom of the tail for subsequent injections.</li>
	<li>Remove the needle and gently apply compression until bleeding stops.</li>
	<li>Return the mouse to the cage, and euthanize the mouse after 3-5 min. 
	NOTE: The mice should be euthanized via isoflurane chamber followed by cervical dislocation. Other methods, such as CO2 inhalation will take up to 5 min and may introduce unnecessary variation of intravenous antibody labeling time.</li>
	<li>Dissect the kidney using scissors, transfer the whole kidney to 1.5 mL of microcentrifuge tubes and leave the samples on ice until further processing. 
	NOTE: Intact whole kidney can be safely stored on ice for a few hours before subsequent processing and digestion.</li>
</ol>
3. Enzymatic digestion of the kidney
<ol>
	<li>Prepare 6 well plates containing 3mL of the digestion solution (RPMI/2% FCS/1 mg/ml Collagenase B) for each mouse, store on ice. 
	NOTE: Stock collagenase B solution at higher concentration (e.g., 20x) can be prepared and stored at or below -20&#176;C. Working solution should be prepared fresh. Please see Table 1 for the recipes for all solution/buffer used in the current protocol.</li>
	<li>Add 300 &#956;L of digestion solution into the 1.5 mL microcentrifuge sample tubes and mince the kidney samples with a straight spring scissor. 
	NOTE: The kidney tissue should be minced into even pieces with sizes no larger than 1.5 mm in diameters. No decapsulation step is required.</li>
	<li>Transfer the minced kidney samples to 6 well plates containing the digestion solution 
	NOTE: 6 well plates are used for convenience only when there are multiple samples to be processed.</li>
	<li>Incubate the samples at 37 &#176;C with gentle rocking (around 60 rpm) for 45 min.</li>
	<li>After digestion, mash the tissue with the plunger flange of a 3 mL syringe directly inside the dish, and transfer into 15 mL conical tubes. 
	NOTE: Usually filtration through a cell strainer is not required at this step because density gradient centrifugation is performed next to purify live lymphocytes.</li>
	<li>Spin down the samples at 500 x g for 5 min at 4 &#176;C.</li>
	<li>Resuspend the pellet in 3 mL of RPMI/10% FCS.</li>
</ol>
4. Density gradient centrifugation to enrich lymphocytes from the digested kidney
<ol>
	<li>Spin down the sample at 500 x g for 5 min at 4 &#176;C.</li>
	<li>Remove the supernatant and resuspend the cell pellet with 5 mL of 44% Percoll/RPMI mix (44% Percoll + 56% RPMI without FBS).</li>
	<li>Put the tip of a 3 mL pipette containing 3 mL of 67% Percoll/PBS (67% Percoll + 33% PBS) directly to the bottom of each tube, and slowly release the solution so that the heavy solution (67% Percoll/PBS) forms a distinct layer at the bottom and the light solution (44% Percoll/RPMI) is raised up. Together, cocktail layers will form. 
	NOTE: For newly arrived density gradient medium from the vendor, add 1/10 volume of sterile 10x PBS to the bottle, mix well and consider the PBS-balanced density gradient medium as 100% solution.</li>
	<li>Spin the samples at 900 x g for 20 min at room temperature with reduced accelerator and brake setting. 
	NOTE: For centrifuges with accelerator and brake settings (on a 1-9 scale (1 means minimum and 9 means maximum)), use 6 for accelerator and 4 for brake.</li>
	<li>Carefully remove the tubes from the centrifuge without disturbing the layers; 
	NOTE: A clear lymphocyte layer is identified between pink color RPMI layer and colorless PBS layer.</li>
	<li>Remove the top layer with a transfer pipette without touching the lymphocyte layer.</li>
	<li>Transfer the lymphocyte layer to a new 15 mL tube, fill the tube with PBS/5% FCS and mix by inverting the tube 4-6x slowly to ensure sufficient mixing. 
	NOTE: This step is important to wash out any residual Percoll.</li>
	<li>Spin down the samples at 500 x g for 5 min at 4 &#176;C.</li>
	<li>Remove the supernatant and re-suspend the cell pellet with 500 &#956;L of complete RPMI medium. The cells are ready for flow cytometry staining.</li>
</ol>
5. Flow cytometry staining and analysis
NOTE: Please follow standard flow cytometry staining protocol for T lymphocytes.
<ol>
	<li>Take around 1 x 106 cells for each FACS staining. 
	NOTE: Use a U-bottom 96 well plate for FACS staining. However, if only a limited number of samples are involved, 5 mL FACS tubes can also be used.</li>
	<li>Spin down the cells at 500 x g for 5 min at 4 &#176;C. Discard the supernatant.</li>
	<li>Resuspend each sample in 50 &#181;L of FACS buffer containing 10 &#181;g/mL of 2.4G2 FcR blocker (an anti-CD16/32 monoclonal antibody to block the interaction between added FACS antibody with FcR). Incubate on ice for 15 min.</li>
	<li>Without further centrifugation, add 50 &#181;L of surface staining antibody mixture for each sample so that the final staining volume is 100 &#181;L/each sample. Incubate on ice for 30 min in the dark. 
	NOTE: Include fluorescently labeled streptavidin (2 &#181;g/mL or follow manufacturer&#8217;s recommendation) in the antibody mixture to label CD8&#945;+ intravascular CD8+ T cells and use fluorescent labeled anti-CD8&#946; antibody to label all CD8 T cells. Antibody mixture is made by adding desired amounts of interested antibodies into the FACS buffer.</li>
	<li>Wash the samples with PBS twice. For each wash, fill the wells of 96 well plate with cold PBS, spin at 500 x g for 5 min at 4 &#176;C and discard the supernatant.</li>
	<li>Resuspend the cells in 100 &#181;L/well of diluted live/death dye. Incubate on ice for 30 min in the dark. 
	NOTE: Even with density gradient medium spin, enzymatic digestion step often induces significant cell death in tissue-resident T cells due to ARTC2.2/P2RX7 signaling15. Live/death dye staining is highly recommended before fixation. Anti-ARTC2 nanobody can be administrated into the mice before euthanasia to prevent cell isolation-induced cell death.</li>
	<li>Wash the samples with FACS buffer twice. For each wash, fill the wells of 96-well plate with cold FACS buffer, spin at 500 x g for 5 min at 4 &#176;C and discard the supernatant.</li>
	<li>Resuspend the cells in 100 &#956;L/well fixing buffer. Incubate at 37 &#176;C for 10 min. 
	NOTE: Fixing buffer is made freshly by diluting formaldehyde with PBS to a final concentration of 2% formaldehyde.</li>
	<li>Wash the samples with FACS buffer twice. For each wash, fill the wells of 96 well plate with cold FACS buffer, spin at 500 x g for 5 min at 4 &#176;C and discard the supernatant.</li>
	<li>Resuspend the cells in 250 &#956;L/well FACS buffer. Seal the plate, store at 4 &#176;C and protect from light until FACS analysis. 
	NOTE: Before FACS analysis, filter the samples through a 70 &#956;M nylon mesh to remove possible cell clusters that may clog the FACS machine.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+emerging+role+of+resident+memory+T+cells+in+protective+immunity+and+inflammatory+disease.">The emerging role of resident memory T cells in protective immunity and inflammatory disease.</a> Nature Medicine. 21, 688-697 (2015).</li><li>Schenkel, J. M., 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=T+cell+memory.+Resident+memory+CD8+T+cells+trigger+protective+innate+and+adaptive+immune+responses.">T cell memory. Resident memory CD8 T cells trigger protective innate and adaptive immune responses.</a> Science. 346, 98-101 (2014).</li><li>Thome, J. J., Farber, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+concepts+in+tissue-resident+T+cells:+lessons+from+humans.">Emerging concepts in tissue-resident T cells: lessons from humans.</a> Trends in Immunology. 36, 428-435 (2015).</li><li>Mackay, L. K., 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=Hobit+and+Blimp1+instruct+a+universal+transcriptional+program+of+tissue+residency+in+lymphocytes.">Hobit and Blimp1 instruct a universal transcriptional program of tissue residency in lymphocytes.</a> Science. 352, 459-463 (2016).</li><li>Mackay, L. 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The authors have no relevant financial disclosures.

As tissue specific immunity is a rapid expanding area of research, accumulating evidence suggest that immune cells, especially lymphocytes population can be identified in almost all organs in adult human or infected or immunized mice. LCMV mouse infection model is a well-established model to study antigen-specific T cell response, effector and memory T cell differentiation including TRM biology across multiple tissues. Here, we described a protocol to analyze CD8+ T cells in the kidney. This protocol is largely adapted from publications focused on TRM12. Presumably due to high level P2RX7 expression and enzymatic digestion induced cell death15, lymphocytes yield from the current protocol is best suited for immediate phenotypic analysis. However, with established protocol to inhibit P2RX7 induced cell death16, it is conceivable that the current protocol can be easily modified (e.g., with the addition of inhibitors to block P2RX7 signaling) to increase the cell survival and be suited for other long-term functional assays. Proper control groups should be included to ensure that P2RX7 inhibitors do not interfere with interested functional assays.
Kidney CD8+ TRM cells are largely generated in response to infection or environmental antigens. We have use intravascular labeling protocol to directly analyze kidney CD8+ T cells isolated from 5-6-week-old na&#239;ve C57BL/6 mice. Consistent with published results17, there is almost no extravascular CD8+ T cells in these &#8220;clean&#8221; mice (Figure 1C). An elegant study has demonstrated that the majority of memory CD8+ T cells identified in non-lymphoid tissues are TRM10. In contrast, using a similar infection system, we often detect a significant population of CD69- cells (most likely represent TEM cells or CD69- TRM cells) even in the extravascular compartment. The discrepancy may be due to the facts that 1) different techniques are used, i.e., microscope vs flow cytometry; 2) early vs late memory time points are focused and 3) CD69 is not a perfect marker to identify TRM. Enzymatic digestion and flow cytometry will significantly under-estimate the total number of TRM cells. However, as a convenient and non-labor-intensive protocol, it is still commonly accepted in most TRM studies.
The gold standard to definitively identify TRM cells is to perform parabiosis experiment. Although the protocol described here provides a convenient way to identify kidney-resident T cells, the results from this protocol only prove that at the time when the mice are euthanized, the T cells are residing outside the blood vessels in the kidney. This protocol alone does not provide any information about the migratory history of T cells.
In addition to infections, any kidney targeting immune responses, including autoimmune responses may induce the differentiation of a significant subset of kidney-resident T cells can be studied using a similar protocol. Further, as described before12, this protocol can be easily modified to use anti-CD45 antibody (to label all hematopoietic cells) to study other kidney-resident immune cells. Together, we demonstrated a relatively convenient way to isolate and analyze CD8+ T cells from the kidney, which can be adapted to various models including infection and autoimmunity.

Tissue-resident memory (TRM) T cells represent one of the most abundant T cell populations in adult human and infected mice. TRM cells provide the first line of immune defense and are critically involved in various physiological and pathological processes1,2,3,4,5. Comparing with circulating T cells, TRM cells carry distinct surface markers with unique transcription programs6,7,8. Expanding our knowledge of TRM biology is the key to understanding T cell responses which is essential for future development of T cell-based vaccines and immunotherapies.
In addition to commonly shared molecular markers of TRMs across all non-lymphoid tissues, accumulating evidence suggest that tissue-specific features are a central component of TRM biology9. Kidney harbors many immune cells including TRM cells after infection and offers a great opportunity to study TRM cell biology in a non-mucosal tissue. Acute LCMV (lymphocytic choriomeningitis virus) infection via intraperitoneal route is a well-established systemic infection model to study antigen-specific T cell responses in mice. The infection is usually resolved in 7-10 days in wild type mice and leave a large number of LCMV-specific memory T cells in a variety of tissues, including the kidney10. P14 TCR (T cell receptor) transgenic mice carry CD8+ T cells recognize one of the immune-dominant epitopes of LCMV presented by MHC-I (class I major histocompatibility complex) molecule H2-Db in C57BL/6 mice. Combining congenically marked P14 T cell adoptive transfer and LCMV infection in mice, CD8+ effector and memory T cells are tracked, including TRM differentiation and homeostasis.
In some barrier tissues, such as intestinal intraepithelial lymphocytes (IEL) compartment and salivary glands, established lymphocyte isolation procedure yields high percentage of TRM cells with minimal blood borne T cell contamination in mouse LCMV model11. However, in non-barrier tissues, such as the kidney, dense vascular network contains a large number of circulating CD8+ T cells. It has been well documented that even successful perfusion cannot efficiently remove all circulating CD8+ T cells. To overcome this technical hurdle, intravascular antibody staining has been established as one of the most commonly used techniques in TRM labs12. In brief, 3-5 minutes before euthanasia, 3 &#181;g/mouse anti-CD8 antibody (to label CD8+ T cells) is delivered intravenously. Intact blood vessel wall prevents rapid diffusion of the antibody within this short period of time (i.e., 3-5 minutes) and only intravascular cells are labeled. Following standard lymphocyte isolation protocol, intravascular versus extravascular cells can be easily distinguished using flow cytometry.
Here, we will describe a standard protocol commonly used in TRM labs to perform intravascular labeling, lymphocyte isolation and flow cytometry analysis of kidney CD8+ T cells using a C57BL/6 mouse that has received CD45.1+ P14 T cells and LCMV infection 30 days before13. Same protocol can be used to study both effector and memory T cells in the kidney.

<table>
	<tbody>
		<tr>
			<td>1.5 ml microcentrifuge tubes</td>
			<td>Fisherbrand</td>
			<td>05-408-130</td>
			<td></td>
		</tr>
		<tr>
			<td>15 ml Conical Tubes</td>
			<td>Corning</td>
			<td>431470</td>
			<td></td>
		</tr>
		<tr>
			<td>3 ml syringe</td>
			<td>BD</td>
			<td>309657</td>
			<td></td>
		</tr>
		<tr>
			<td>37C incubator</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Biotin a-CD8a antibody(Clone 53-6.7)</td>
			<td>Tonbo Biosciences</td>
			<td>30-0081-U025</td>
			<td></td>
		</tr>
		<tr>
			<td>Calf Serum</td>
			<td>GE Healthcare Life Sciences</td>
			<td>SH30073.03</td>
			<td></td>
		</tr>
		<tr>
			<td>Collegenase B</td>
			<td>Millipore Sigma</td>
			<td>11088807001</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable Graduated Transfer Pipettes</td>
			<td>Fisherbrand</td>
			<td>13-711-9AM</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin Syringe</td>
			<td>BD</td>
			<td>329424</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Dissecting Spring Scissors</td>
			<td>Roboz Surgical</td>
			<td>RS 5692</td>
			<td></td>
		</tr>
		<tr>
			<td>Mojosort Mouse CD8 Na&#239;ve T Cells Isolation Kit</td>
			<td>Biolegend</td>
			<td>480043</td>
			<td></td>
		</tr>
		<tr>
			<td>overhead heat lamp</td>
			<td>Amazon</td>
			<td>B00333PZZG</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Percoll</td>
			<td>GE Healthcare Life Sciences</td>
			<td>17089101</td>
			<td></td>
		</tr>
		<tr>
			<td>rocker</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI</td>
			<td>GE Healthcare Life Sciences</td>
			<td>SH30096.01</td>
			<td></td>
		</tr>
		<tr>
			<td>Solid Brass Mouse Restrainer</td>
			<td>Braintree Scientific, Inc</td>
			<td>SAI-MBR</td>
			<td></td>
		</tr>
		<tr>
			<td>Swing Bucket Centrifuge with refrigerator</td>
			<td>Thermofisher</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Culture 6-well plate</td>
			<td>Corning</td>
			<td>3516</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation of mouse kidney-resident cd8+ t cells for flow cytometry analysis

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the key steps to investigate TRMs. There are slight variations in lymphocyte isolation protocols for different organs. Kidney is an essential non-lymphoid organ with numerous immune cell infiltration especially after pathogen exposure or autoimmune activation. In recent years, multiple labs including our own have started characterizing kidney resident CD8+ T cells in various physiological and pathological settings in both mouse and human. Due to the abundance of T lymphocytes, kidney represents an attractive model organ to study TRMs in non-mucosal or non-barrier tissues. Here, we will describe a protocol commonly used in TRM-focused labs to isolate CD8+ T cells from mouse kidneys following systemic viral infection. Briefly, using an acute lymphocytic choriomeningitis virus (LCMV) infection model in C57BL/6 mice, we demonstrate intravascular CD8+ T cell labeling, enzymatic digestion, and density gradient centrifugation to isolate and enrich lymphocytes from mouse kidneys to make samples ready for the subsequent flow cytometry analysis.

The protocol described here is summarized in a flow chart (Figure 1A). At day 30 post LCMV infection, we performed intravascular labeling of CD8+ T cells. 5 minutes later, both kidneys of the animal were dissected, minced and subjected to collagenase digestion. Lymphocytes were further purified from the digested samples via Percoll centrifugation and analyzed by flow cytometry. As shown in Figure 1B, even after density centrifugation-mediated lymphocyte enrichment, it was very common to see a large portion of non-lymphocytes in the final product. However, after gating on live lymphocytes, it was easy to identify CD8+ T lymphocytes. We could distinguish intravascular vs extravascular CD8+ T cells. Due to highly correlative expression pattern of CD8&#945; and CD8&#946; on CD8+ T cells, it is expected that intravascular CD8+ T cells exhibit a diagonal pattern on CD8&#945;-CD8&#946; FACS profile. As expected, only extravascular CD8+ T cell efficiently acquired TRM phenotypes, such as upregulation of CD69 and downregulation of Ly6C (Figure 1B). In our experiments, we were interested in donor P14 T cells with congenic marker CD45.1. Using the same protocol, endogenous host derived CD8+ T cells can be examined as well. In contrast to infected mice, the vast majority of CD8+ T cells isolated from young na&#239;ve mice housed in SPF facility were labeled with i.v. CD8&#945; antibody and, therefore, belonged to the intravascular compartment (Figure 1C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61559/61559fig01.jpg" /> 
Figure 1: Representative results. 
(A) Flowchart of the protocol. (B) At day 30 post infection, kidney lymphocytes were analyzed by flow cytometry. Representative FACS profiles and gating strategy are shown. The numbers indicate the percentage of gated subsets in their parental population. (C) Representative FACS profile of kidney CD8+ T cells isolated from a na&#239;ve mouse. <a href="https://www.jove.com/files/ftp_upload/61559/61559fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Recipe</td>
			<td>NOTE</td>
		</tr>
		<tr>
			<td>100% Percoll</td>
			<td>9 volumes of Percoll + 1 volume of 10xPBS</td>
			<td></td>
		</tr>
		<tr>
			<td>44% Percoll/RPMI</td>
			<td>44% Vol of 100% Percoll + 56% Vol serum free RPMI</td>
			<td>Made fresh from 100% Percoll</td>
		</tr>
		<tr>
			<td>67% Percoll/PBS</td>
			<td>67% Vol of 100% Percoll + 33% Vol PBS</td>
			<td>Made fresh from 100% Percoll</td>
		</tr>
		<tr>
			<td>Digestion buffer</td>
			<td>RPMI + 2% FCS + 1mg/ml collagenase B</td>
			<td>Made fresh from collagenase stock</td>
		</tr>
		<tr>
			<td>PBS/5% FCS washing buffer</td>
			<td>PBS + 5% FCS</td>
			<td></td>
		</tr>
		<tr>
			<td>FACS buffer</td>
			<td>PBS + 2% FCS + 0.02% NaN3</td>
			<td>Stored at 4&#176;C</td>
		</tr>
		<tr>
			<td>Fixing buffer</td>
			<td>PBS + 2% formaldehyde</td>
			<td>Stored at RT and protected from light</td>
		</tr>
	</tbody>
</table>
Table 1: List of recipes for all solution and buffer used in current protocol.

Watch this Scientific Journal Video about Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis at JoVE.com

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Following viral infection, kidney harbors a relatively large number of CD8+ T cells and offers an opportunity to study non-mucosal TRM cells. Here, we describe a protocol to isolate mouse kidney lymphocytes for flow cytometry analysis.

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the ...

isolation-mouse-kidney-resident-cd8-t-cells-for-flow-cytometry

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

6.5K Views.  University of Texas Health Science Center at San Antonio. Following viral infection, kidney harbors a relatively large number of CD8+ T cells and offers an opportunity to study non-mucosal TRM cells. Here, we describe a protocol to isolate mouse kidney lymphocytes for flow cytometry analysis.

Video: Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

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 ...

Whole-animal Imaging and Flow Cytometric Techniques for Analysis of Antigen-specific CD8+ T Cell Responses after Nanoparticle Vaccination

Traditional vaccine adjuvants, such as alum, elicit suboptimal CD8+ T cell responses. To address this major challenge in vaccine development, various nanoparticle systems have been engineered to mimic features of pathogens to improve antigen delivery to draining lymph nodes and increase antigen uptake by antigen-presenting cells, leading to new vaccine formulations optimized for induction of antigen-specific CD8+ T cell responses. In this article, we describe the synthesis of a &#8220;pathogen-mimicking&#8221; nanoparticle system, termed interbilayer-crosslinked multilamellar vesicles (ICMVs) that can serve as an effective vaccine carrier for co-delivery of subunit antigens and immunostimulatory agents and elicitation of potent cytotoxic CD8+ T lymphocyte (CTL) responses. We describe methods for characterizing hydrodynamic size and surface charge of vaccine nanoparticles with dynamic light scattering and zeta potential analyzer and present a confocal microscopy-based procedure to analyze nanoparticle-mediated antigen delivery to draining lymph nodes. Furthermore, we show a new bioluminescence whole-animal imaging technique utilizing adoptive transfer of luciferase-expressing, antigen-specific CD8+ T cells into recipient mice, followed by nanoparticle vaccination, which permits non-invasive interrogation of expansion and trafficking patterns of CTLs in real time. We also describe tetramer staining and flow cytometric analysis of peripheral blood mononuclear cells for longitudinal quantification of endogenous T cell responses in mice vaccinated with nanoparticles.

We describe whole-animal imaging and flow cytometry-based techniques for monitoring expansion of antigen-specific CD8+ T cells in response to immunization with nanoparticles in a murine model of vaccination.

Traditional vaccine adjuvants, such as alum, elicit suboptimal CD8+ T cell responses. To address this major challenge in vaccine development, various ...

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

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

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

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

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method enables researchers to better understand the characteristics of a single cell population without the influence of other cells. Compared to other methods of cell enrichment, such as magnetic-activated cell sorting (MCS), FACS is more flexible and accurate for cell separation due to the ability of phenotype detection by flow cytometry. In addition, FACS is usually capable of separating multiple cell populations simultaneously, which improves the efficiency and diversity of experiments. Although FACS has some limitations, it has been broadly used to purify cells for functional studies in both in vitro and in vivo settings. Here we report a protocol using fluorescence-activated cell sorting to isolate a very rare population of immune cells, plasmacytoid dendritic cells (pDC), with high purity from the bone marrow of lupus-prone mice for in vitro functional studies of pDC.

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of pDC.

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method ...

Isolation and Flow Cytometric Analysis of Human Endocervical Gamma Delta T Cells

The female reproductive tract (FRT) mucosal immune system serves as the first line of defense. Better knowledge of the genital mucosa is therefore essential for understanding pathogenicity of different pathogens including HIV. Gamma delta (GD) T cells are the prototype of &#39;unconventional&#39; T cells and represent a relatively small subset of T cells defined by their expression of heterodimeric T-cell receptors (TCRs) composed of gamma and delta chains. This sets them apart from the classical and much better known CD4+ helper T cells and CD8+ cytotoxic T cells that are defined by alpha-beta TCRs. GD T cells often show tissue-specific localization and are enriched in epithelium. GD T cells orchestrate immune responses in inflammation, tumor surveillance, infectious disease, and autoimmunity.
Here, we present a method to reproducibly isolate and analyze human endocervical intraepithelial GD T lymphocytes. We have used endocervical cytobrush samples from women participating in the Women&#39;s Interagency HIV Infection Study (WIHS). Knowledge about GD T cells interactions during conditions in which there is an insult to the vaginal mucosal could be applied to any clinical study in which mucosal vulnerability is addressed, including the development of vaginal microbicides.In addition, knowledge about mucosal GD T cell responses has potential for application of GD T cell-based immune therapy in treating infectious diseases.

We describe here an isolation method to obtain human endocervical intraepithelial lymphocytes for the analysis of intraepithelial gamma delta T cells. This protocol can be extended for the purification of endocervical gamma delta T cells by magnetic beads or by cell sorting.

The female reproductive tract (FRT) mucosal immune system serves as the first line of defense. Better knowledge of the genital mucosa is therefore ...

Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal of immunosuppression. The mixed leukocyte reaction (MLR), limiting dilution assays, and trans-vivo delayed-type hypersensitivity (DTH) assay have all been applied to this question, but these methods have limited predictive ability and/or significant practical limitations that reduce their usefulness. Imaging flow cytometry is a technique that combines the multiparametric quantitative powers of flow cytometry with the imaging capabilities of fluorescent microscopy. We recently made use of an imaging flow cytometry approach to define the proportion of recipient T cells capable of forming mature immune synapses with donor antigen-presenting cells (APCs). Using a well-characterized mouse heart transplant model, we have shown that the frequency of in vitro immune synapses among T-APC membrane contact events strongly predicted allograft outcome in rejection, tolerance, and a situation where transplant survival depends on induced regulatory T cells. The frequency of T-APC contacts increased with T cells from mice during acute rejection and decreased with T cells from mice rendered unresponsive to alloantigen. The addition of regulatory T cells to the in vitro system reduced prolonged T-APC contacts. Critically, this effect was also seen with human polyclonally expanded, naturally occurring regulatory T cells, which are known to control the rejection of human tissues in humanized mouse models. Further development of this approach may allow for a deeper characterization of the alloreactive T-cell compartment in transplant recipients. In the future, further development and evaluation of this method using human cells may form the basis for assays used to select patients for immunosuppression minimization, and it can be used to measure the impact of tolerogenic therapies in the clinic.

This paper describes a method for measuring alloreactivity in a mixed population of T cells using imaging flow cytometry.

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...

A DNA/Ki67-Based Flow Cytometry Assay for Cell Cycle Analysis of Antigen-Specific CD8 T Cells in Vaccinated Mice

The cell cycle of antigen-specific T cells in vivo has been examined by using a few methods, all of which possess some limitations. Bromodeoxyuridine (BrdU) marks cells that are in or recently completed S-phase, and carboxyfluorescein succinimidyl ester (CFSE) detects daughter cells after division. However, these dyes do not allow identification of the cell cycle phase at the time of analysis. An alternative approach is to exploit Ki67, a marker that is highly expressed by cells in all phases of the cell cycle except the quiescent phase G0. Unfortunately, Ki67 does not allow further differentiation as it does not separate cells in S-phase that are committed to mitosis from those in G1 that can remain in this phase, proceed into cycling, or move into G0.
Here, we describe a flow cytometric method for capturing a &#34;snapshot&#34; of T cells in different cell cycle phases in mouse secondary lymphoid organs. The method combines Ki67 and DNA staining with major histocompatibility complex (MHC)-peptide-multimer staining and an innovative gating strategy, allowing us to successfully differentiate between antigen-specific CD8 T cells in G0, in G1 and in S-G2/M phases of the cell cycle in the spleen&#160;and draining lymph nodes of mice after vaccination with viral vectors carrying the model antigen gag of human immunodeficiency virus (HIV)-1.
Critical steps of the method were the choice of the DNA dye and the gating strategy to increase the assay sensitivity and to include highly activated/proliferating antigen-specific T cells that would have been missed by current criteria of analysis. The DNA dye, Hoechst 33342, enabled us to obtain a high-quality discrimination&#160;of the G0/G1 and G2/M DNA peaks, while preserving membrane and intracellular staining. The method has great potential to increase knowledge about T cell response in vivo and to improve immuno-monitoring analysis.

Clonal expansion is a key feature of antigen-specific T cell response. However, the cell cycle of antigen-responding T cells has been poorly investigated, partly because of technical limitations. We describe a flow cytometric method to analyze clonally expanding antigen-specific CD8 T cells in spleen and lymph nodes of vaccinated mice.

The cell cycle of antigen-specific T cells in vivo has been examined by using a few methods, all of which possess some limitations. Bromodeoxyuridine ...

Isolation of Mouse Kidney-Resident CD8⁺ T cells for Flow Cytometry Analysis

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Chapters in this video

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Whole-animal Imaging and Flow Cytometric Techniques for Analysis of Antigen-specific CD8+ T Cell Responses after Nanoparticle Vaccination

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

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Isolation and Flow Cytometric Analysis of Human Endocervical Gamma Delta T Cells

Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

A DNA/Ki67-Based Flow Cytometry Assay for Cell Cycle Analysis of Antigen-Specific CD8 T Cells in Vaccinated Mice