The authors would like to thank the GI and Liver Innate Immune Programs for monetary support of this project. B.A.J.T. is also funded by R01 AI121209.

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Colorado Anschutz Medical Campus.
1.&#160;Preparation of the Materials
<ol>
	<li>Make a 5 mg/mL solution of DNase I in phosphate-buffered saline (PBS).</li>
	<li>Make a digestion mixture by adding 5,000 U/mL of collagenase IV to Click&#8217;s EHAA media.</li>
	<li>Warm the digestion mixture at 37 &#176;C for 30 min prior to use.</li>
	<li>Make an isolation buffer by adding 4.8% bovine serum albumin (BSA) and 2 mM ethylenediaminetetraacetic acid (EDTA) to Hanks&#8217; balanced salt solution (HBSS).</li>
	<li>Make a red blood cell (RBC) lysis buffer by adding 100 mM ammonium chloride, 10 mM KHCO3, and 0.1 mM EDTA to distilled H2O.</li>
</ol>
2. Preparation of a Single-cell Suspension from a Mouse Liver
<ol>
	<li>Euthanize the mouse with CO2 and cervical dislocation.</li>
	<li>Spray down the mouse with 70% ethanol to wet its fur. Pin the mouse&#8217;s feet to a dissection board.</li>
	<li>Using dissection scissors to cut the skin about 1 cm above the anus, being careful to cut only through the skin (about 1 mm). Pull the skin away from the body with toothed forceps and insert the scissors between the skin and peritoneum. Open the scissors to separate the skin from the peritoneum and, then, cut the skin from the incision to the neck.</li>
	<li>Pin the skin to the dissection board using one pin under each arm and above each leg. Pull the peritoneal sac up and cut upwards toward the neck. Grab the lobes of the liver and cut just below the sternum. 
	Note: Care should be taken if any of the liver will be used for immunohistochemistry (IHC).</li>
	<li>Cut around the liver and remove the liver from the mouse and place it in 4 mL of Click&#8217;s EHAA media.</li>
	<li>Using a scalpel, cut the liver in ~1 mm diameter pieces.</li>
	<li>Add 500 &#181;L of the digestion mixture and 500 &#181;L of the DNase I (2 mg/mL) to the liver.</li>
	<li>Incubate the liver for 30 min at 37 &#176;C. After 15 min, mix the liquid using a 5 mL pipette.</li>
	<li>After 30 min of incubation, transfer the digested sample through a 100 &#181;m strainer to a 50 mL conical tube.</li>
	<li>Gently push the remaining pieces through the filter with the plunger of a 1-mL syringe.</li>
	<li>Wash the filter with 5 mL of isolation buffer and gently push the tissue through the strainer with the back of a plunger from a 1 mL syringe. Repeat this until the filter is washed with 25 mL of isolation buffer.</li>
	<li>Centrifuge the cells at 400 x g for 5 min. Carefully aspirate off the supernatant.</li>
	<li>Resuspend the pellet with 4 mL of RBC lysis buffer. Incubate the cells at room temperature for 5 min.</li>
	<li>Wash the cells with 10 mL of isolation buffer and centrifuge at 400 x g for 5 min.</li>
	<li>Count the cells on a hemocytometer to determine the full liver count.</li>
	<li>Resuspend cells in 5 mL of 20% iodixanol and layer them with 1 mL of PBS.</li>
	<li>Centrifuge the cells at 300 x g for 15 min without a brake.</li>
	<li>Remove the layer between the PBS and the iodixanol and place them, through a 100 &#181;m filter, into a new 50 mL conical tube.</li>
	<li>Wash the cells with 10 mL of isolation buffer and centrifuge at 400 x g for 5 min.</li>
	<li>Discard the supernatant and resuspend the cells in 500 &#181;L of PBS with 2% fetal bovine serum (FBS).</li>
</ol>
3.&#160;Flow Cytometric Analysis of Single Cells from the Liver
<ol>
	<li>Count the cells using a hemocytometer and microscope, using trypan blue exclusion to measure viable cells. Add 10 &#181;L of the cells to 10 &#181;L of trypan blue and immediately place them on a hemocytometer and count the live cells (not blue) under a microscope. Then, calculate the number of cells per microliter.</li>
	<li>Aliquot approximately 5 million of the remaining nonparenchymal cells into a single well of a 96-well plate.</li>
	<li>Centrifuge the cells at 400 x g for 5 min.</li>
	<li>Discard the supernatant and resuspend the cells in 90 &#181;L of PBS with 2% FBS.</li>
	<li>Add anti-CD45 (1:200), anti-CD146 (1:200), anti-CD31 (1:200), and PDPN (1:200) diluted in 10 &#181;L of 10x 2.4G2 or anti-CD16/32 (1:200). 
	NOTE: No Fc block (2.4G2) was used when anti-CD16/32-labeled antibody was used.</li>
	<li>To determine where positive and negative gates should be set, include a fluorescence minus one (FMO) stain for each color and an isotype control antibody.</li>
	<li>To determine live versus dead cells, stain with a viability marker (e.g., ghost red 780). Incubate the cells at 4 &#176;C for 30 min.</li>
	<li>Wash the cells with 100 &#181;L of PBS with 2% FBS.</li>
	<li>Use a small aliquot of cells to adjust the laser and compensation settings on the flow cytometer. Stain the cells with an antibody to each individual fluorophore and one without any antibody. 
	NOTE: Depending on the flow cytometer being used, a compensation matrix should be established to remove spectral overlap.</li>
	<li>Place the sample tube onto the cytometer probe and collect and record all events.</li>
</ol>
4.&#160;Data Analysis
<ol>
	<li>Looking at side-scatter area vs. forward-scatter area, gate on &#8220;live&#8221; cells based on size and granularity and viability marker dye.</li>
	<li>Next, using CD45 Brilliant Violet 510 and CD31 PerCp Cy5.5, gate on the CD45- CD31+ cells using the isotype controls and FMO to determine positive and negative populations.</li>
	<li>Lastly, using CD146 v450 or CD16/32 FITC and PDPN APC, take the CD146- PDPN+ or CD16/32- PDPN+ cells, again using isotype controls and FMO, to determine positive and negative populations. These cells are the LECs.</li>
</ol>

<ol>
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N., 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=Lymph+node-resident+lymphatic+endothelial+cells+mediate+peripheral+tolerance+via+Aire-independent+direct+antigen+presentation.">Lymph node-resident lymphatic endothelial cells mediate peripheral tolerance via Aire-independent direct antigen presentation.</a> Journal of Experimental Medicine. 207 (4), 681-688 (2010).</li><li>Cohen, J. N., 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=Tolerogenic+properties+of+lymphatic+endothelial+cells+are+controlled+by+the+lymph+node+microenvironment.">Tolerogenic properties of lymphatic endothelial cells are controlled by the lymph node microenvironment.</a> PLoS One. 9 (2), e87740 (2014).</li><li>Rouhani, S. 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W., 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=VEGF-C+promotes+immune+tolerance+in+B16+melanomas+and+cross-presentation+of+tumor+antigen+by+lymph+node+lymphatics.">VEGF-C promotes immune tolerance in B16 melanomas and cross-presentation of tumor antigen by lymph node lymphatics.</a> Cell Reports. 1 (3), 191-199 (2012).</li><li>Lund, A. W., 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=Lymphatic+vessels+regulate+immune+microenvironments+in+human+and+murine+melanoma.">Lymphatic vessels regulate immune microenvironments in human and murine melanoma.</a> Journal of Clinical Investigation. 126 (9), 3389-3402 (2016).</li><li>Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunomodulatory+roles+of+lymphatic+vessels+in+cancer+progression.">Immunomodulatory roles of lymphatic vessels in cancer progression.</a> Cancer Immunology Research. 2 (8), 701-707 (2014).</li><li>Dietrich, T., 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=Cutting+edge:+lymphatic+vessels,+not+blood+vessels,+primarily+mediate+immune+rejections+after+transplantation.">Cutting edge: lymphatic vessels, not blood vessels, primarily mediate immune rejections after transplantation.</a> Journal of Immunology. 184 (2), 535-539 (2010).</li><li>Kedl, R., 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=Migratory+Dendritic+Cells+acquire+archived+antigen+from+Lymphatic+Endothelial+Cells+for+antigen+presentation+during+lymph+node+contraction.">Migratory Dendritic Cells acquire archived antigen from Lymphatic Endothelial Cells for antigen presentation during lymph node contraction.</a> Nature Communications. 8, 2034 (2017).</li><li>Kedl, R. 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The authors have nothing to disclose.

The overall importance of LECs in immune homeostasis and regulation has recently come to light25. Much of the published lymphatic literature focuses on skin and lymph nodes; however, lymphatics are found throughout the body26 and, thus, our understanding of their importance in different organs is needed. Here we show a method in which LECs in the liver can be studied on a cell-by-cell basis to better understand their concurrent expression of different surface markers, cytokines, chemokines, and intracellular proteins such as transcription factors. This method will be useful for future studies to assess the phenotype and function of LECs in the liver during health and disease.
One of the hurdles for the identification of LECs in the liver is their relatively low frequency compared to other cell types. Hepatocytes make up about 80% of the liver, and removing these cells using density gradient (iodixanol) before running the liver on a flow cytometer requires less time and, thus, provides better viability. In order to distinguish the population of LYVE1+ LECs in the liver from LYVE1+ LSECs, we used the markers CD16/32 and CD146 found on LSECs and with low to no expression by LECs. This, coupled with the lack or low expression of PDPN by any other endothelial cell in the liver, allowed the validation by flow cytometry that the population we identified were LECs. Indeed, downstream transcriptional analysis confirmed that this gating strategy produced LECs (Figure 3C).
The isolation of LECs from the lymph node is best done using collagenase I-II-and-dispase; however, we found that, while this method does extract LECs from the liver, a liver digestion protocol using collagenase IV provides a better downstream analysis using flow cytometry. Using a mechanical disruption of the liver allows the collagenase more surface area to better interact with the extracellular matrix-associated cells, like LECs, and using Click&#39;s EHAA media without FBS allows for the digestion of the liver to occur in only 30 min. This decreased time maintains LEC viability for downstream assays like flow cytometry or flow sorting. Indeed, we were able to recover enough viable cells to visualize LECs by flow cytometry and flow sorting for downstream transcriptional analysis.
Combined, the separation of hepatocytes from the non-parenchymal cells, the use of collagenase type IV, and the clarification and demonstration of markers specific to LECs in the liver and markers specific to other endothelial cell populations, such as CD16/32 and CD146, allowed the proper identification of LECs in the liver. These methods fill a significant gap in the literature about how LECs in the liver can be identified by flow cytometry, especially since the liver contains a number of other cells that express markers known to be unique to LECs in the lymph node (prox1 and vegfr3). Therefore, these methods will lead to downstream studies regarding liver LEC function. Additionally, this method can be modified for other tissues in order to better evaluate tissue-specific LEC markers and subsets.

The role of lymphatic vessels and LECs in the liver is not well understood. While lymphatic vessels are found within the portal triad of the liver1 and expand during disease2, very little is understood regarding the function and phenotype of LECs within the liver. With the discovery of markers that are found primarily on LECs3, the importance of these cells within different tissue niches in homeostasis and disease will fill a significant gap in our understanding. LECs have a major role in maintaining peripheral tolerance in the lymph node and in metastatic tumors by interacting directly with T cells4,5,6,7,8,9,10,11,12,13. LECs in the lymph node can promote protective immunity via their interactions with migratory dendritic cells14,15,16. Therefore, there are multiple roles for LECs which may be specific to the tissues and interactions in which they are present. However, very little is understood about how LECs interact with immune cells in the tissue or how LECs function in different organ systems; thus, evaluating LECs on a per cell basis within the liver or other organs may lead to advances in how LECs program tissue-specific immunity. While much of the literature that focuses on LECs in the liver uses microscopy to visualize LECs using one or two markers and morphology17, very little has been done to specifically evaluate LECs on a cell by cell basis using flow cytometry, though one study did evaluate differences between liver sinusoidal endothelial cells (LSECs) and LECs18. Being able to analyze LEC populations in the liver by flow cytometry allows for the in-depth study of LEC phenotype during normal homeostasis or disease.
To evaluate LECs by flow cytometry, multiple surface markers are needed. Typically, LECs are visualized by the expression of prospero-related homeobox 1 (Prox-1), lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1) or vascular endothelial growth factor receptor 3 (VEGFR3) using microscopy. However, in the liver, the expression of these markers is not restricted to LECs. Prox-1 is widely expressed by hepatocytes during liver development, regeneration, and injury19, and LYVE1 and VEGFR3 are expressed by the liver sinusoidal endothelial cells18. In the lymph node, LECs are identified using flow cytometry as clusters of differentiation (CD) CD45-, CD31+, and podoplanin+ (PDPN)16. However, this approach is too minimal to isolate LECs in the liver since CD45- CD31+ cells will capture endothelial cells, and the predominant population of vascular endothelial cells in the liver are LSECs. Thus, other markers are needed to distinguish the rare LEC population from the abundant LSEC population. Both CD16/32 (expressed by mature LSECs18) and CD146 (a common vascular endothelial cell marker that is predominately expressed within the liver sinusoids by liver sinusoidal endothelial cells20 with little to no expression by lymphatic endothelial cells21) were candidate markers.
Therefore, we optimized a method for isolating and visualizing LECs in the liver using the above markers, CD45, CD31, CD146, CD16/32, and PDPN for flow cytometry. We describe the use of collagenase IV, DNase 1, and mechanical separation for liver tissue digestion into a single-cell suspension. We also describe the use of iodixanol density gradient for the isolation of non-parenchymal cells (NPC) and to eliminate cellular debris. Finally, using multiple markers, we determine the optimal flow cytometry gating strategy to identify LECs from the liver with PDPN as the predominant marker.

<table>
	<tbody>
		<tr>
			<td>Clicks/EHAA media</td>
			<td>Irvine Scientific</td>
			<td>9195</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase IV</td>
			<td>Worthington Biochemical corporation</td>
			<td>LS004188</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I</td>
			<td>Worthington Biochemical corporation</td>
			<td>LS002145</td>
			<td>Deoxyribonuclease 1</td>
		</tr>
		<tr>
			<td>OptiPrep</td>
			<td>Sigma Aldrich</td>
			<td>D1556</td>
			<td>Density Gradient Medium</td>
		</tr>
		<tr>
			<td>V450 anti mouse CD146(clone ME-9F1</td>
			<td>BD biosciences</td>
			<td>562232</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti mouse CD146 (clone ME-9F1</td>
			<td>Biolegend</td>
			<td>134706</td>
			<td>Fluorescein isothiocyanate (FITC)</td>
		</tr>
		<tr>
			<td>Pacific Blue anti mouse CD31(clone 390)</td>
			<td>Biolegend</td>
			<td>102422</td>
			<td></td>
		</tr>
		<tr>
			<td>PerCp/Cy5.5 anti mouse CD31( clone 390)</td>
			<td>Biolegend</td>
			<td>102420</td>
			<td>Peridinin-chlorophyll proteins-Cyanine 5.5 (PerCP-Cy5.5)</td>
		</tr>
		<tr>
			<td>APC anti mouse PDPN (clone 8.1.1)</td>
			<td>Biolegend</td>
			<td>127410</td>
			<td>Allophycocyanin (APC), podoplanin (PDPN)</td>
		</tr>
		<tr>
			<td>APC/Cy7 anti mouse CD45 (clone 30-F11)</td>
			<td>Biolegend</td>
			<td>103116</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 510 anti mouse CD45 (clone 30-F11</td>
			<td>Biolegend</td>
			<td>103138</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti mouse CD16/32 (clone 93)</td>
			<td>Biolegend</td>
			<td>101306</td>
			<td>Fluorescein isothiocyanate (FITC)</td>
		</tr>
		<tr>
			<td>PerCp/Cy5.5 anti mouse CD16/32( clone 93)</td>
			<td>Biolegend</td>
			<td>101324</td>
			<td>Peridinin-chlorophyll proteins-Cyanine 5.5 (PerCP-Cy5.5)</td>
		</tr>
		<tr>
			<td>ghost red 780 viability dye</td>
			<td>TONBO biosceinces</td>
			<td>3-0865-T100</td>
			<td></td>
		</tr>
		<tr>
			<td>APC syrian hamster IgG (clone SHG-1)</td>
			<td>Biolegened</td>
			<td>402102</td>
			<td></td>
		</tr>
		<tr>
			<td>PerCp/Cy5.5 rat IgG2a (clone RTK2758)</td>
			<td>Biolegend</td>
			<td>400531</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC rat IgG2 (clone eBR2a)</td>
			<td>ebioscience</td>
			<td>1-4321-80</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti mouse LYVE1 (clone 223322)</td>
			<td>R&#38;D systems</td>
			<td>FAB2125A</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-mouse Cytokeratin(clone EPR17078)</td>
			<td>abcam</td>
			<td>ab181598</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-mouse F4/80 (clone Cl:A3-1)</td>
			<td>Bio-rad</td>
			<td>MCA497</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA (fraction V)</td>
			<td>Fischer</td>
			<td>BP1600-100</td>
			<td>Bovine Serum Albumin (BSA)</td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Jackson Immunoresearch</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Serum</td>
			<td>Jackson Immunoresearch</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>VWR</td>
			<td>E177</td>
			<td>Ethylenediaminetetraacetic acid (EDTA) -for RBC lysis buffer</td>
		</tr>
		<tr>
			<td>Ammonium Chloride</td>
			<td>Fischer</td>
			<td>A687-500</td>
			<td>for RBC Lysis buffer</td>
		</tr>
		<tr>
			<td>Potassium Bicarbonate</td>
			<td>Fischer</td>
			<td>P184-500</td>
			<td>for RBC Lysis buffer</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Feather</td>
			<td>2975#21</td>
			<td></td>
		</tr>
		<tr>
			<td>100um cell strainer</td>
			<td>Fischer</td>
			<td>22363549</td>
			<td></td>
		</tr>
		<tr>
			<td>2.4G2</td>
			<td>in house/ATCC</td>
			<td>ATCC HB-197</td>
			<td>FC block to inhibit non-specific binding to Fc gamma + cells -made from hybridoma</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Corning</td>
			<td>21-040-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks Balanced Salt Solution (HBSS)</td>
			<td>Gibco</td>
			<td>14185-052</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Atlanta biologicals</td>
			<td>S11550</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well plate</td>
			<td>Corning</td>
			<td>3788</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plate</td>
			<td>Corning</td>
			<td>3506</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml conical</td>
			<td>Truline</td>
			<td>TR2004</td>
			<td></td>
		</tr>
		<tr>
			<td>15 ml conical</td>
			<td>Falcon</td>
			<td>352196</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml Pipete tip</td>
			<td>USA scientific</td>
			<td>1111-2721</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;l pipete tip</td>
			<td>USA scientific</td>
			<td>1110-1700</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;l pipete tip</td>
			<td>USA scientific</td>
			<td>1111-3700</td>
			<td></td>
		</tr>
		<tr>
			<td>seriological 10ml pipete</td>
			<td>greiner bio-one</td>
			<td>607107</td>
			<td></td>
		</tr>
		<tr>
			<td>seriological 5ml pipete</td>
			<td>greiner bio-one</td>
			<td>606107</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell incubator</td>
			<td>Fischer</td>
			<td></td>
			<td>Heracell 160i</td>
		</tr>
		<tr>
			<td>BD FacsCanto II flow cytometer</td>
			<td>BD biosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Clinical Centrifuge</td>
			<td>Beckman coulter</td>
			<td></td>
			<td>model X-14R</td>
		</tr>
	</tbody>
</table>

digestion of the murine liver for a flow cytometric analysis of lymphatic endothelial cells

Within the liver, lymphatic vessels are found within the portal triad, and their described function is to remove interstitial fluid from the liver to the lymph nodes where cellular debris and antigens can be surveyed. We are very interested in understanding how the lymphatic vasculature might be involved in inflammation and immune cell function within the liver. However, very little has been published establishing digestion protocols for the isolation of lymphatic endothelial cells (LECs) from the liver or specific markers that can be used to evaluate liver LECs on a per cell basis. Therefore, we optimized a method for the digestion and staining of the liver in order to evaluate the LEC population in the liver. We are confident that the method outlined here will be useful for the identification and isolation of LECs from the liver and will strengthen our understanding of how LECs respond to the liver microenvironment.

Studies analyzing liver lymphatics have primarily used immunohistochemistry to quantitate the frequency and diameter of lymphatic vessels in the liver. However, this method does not allow for the evaluation of LECs on a cell-by-cell basis or for expression of multiple markers, cytokines, chemokines, or transcription factors. Therefore, we asked whether liver LECs could be isolated from the liver and evaluated using flow cytometry. Previous work isolating lymph node LECs was performed using Liberase DL (collagenase I-II-and-dispase) and DNase 1 combined with the mechanical separation of the lymph node tissue with needles14,16. Therefore, the same digestion protocol on liver tissue was used, or collagenase IV and DNase 1 was used as previously described, for immune cell isolation from the liver22 and followed the digestion with a density gradient separation (iodixanol) step to remove hepatocytes and increase the frequency of other cell types within the liver (Figure 1A). Following liver digestion and centrifugation with the density gradient, the cells were stained with CD45, CD31, PDPN, and CD16/32. CD16/32 has been described to be expressed by mature LSEC populations, but not LEC populations18. Thus, LECs were gated as CD45-, CD31+, CD16/32-, PDPN+ cells, while LSECs were gated as CD45-, CD31+, CD16/32+ and PDPN- (Figure 1A). Gates were set on live cells (ghost red negative) and based on isotype controls and FMO staining (Figure 1A). LYVE-1 APC on the LEC population was also confirmed (Figure 1B). Both methods allowed the visualization of the LEC population within the liver; however, the staining profile of the cells was visually better when using collagenase IV and DNase 1 than collagenase I-II-and-dispase (Figure 1C). Therefore, all future manipulations were performed using collagenase IV and DNase 1, as described in the protocol. On average, we obtained approximately 1,300 LECs per gram of liver tissue or 2,200 LECs per naive liver, using this digestion method.
To optimize the gating strategy and combination of fluorophores and to eliminate contamination of LSECs, both CD16/32 and CD146 were used.&#160;CD16/32 is Fc gamma receptors II and III and is expressed on mature LSECs but not on LECs18. CD16/32 could distinguish the PDPN+ CD16/32- cells from the PDPN- CD16/32+ cells, especially when using PDPN conjugated to APC or PE and CD16/32 conjugated to FITC (Figure 1A), but less well when PDPN was conjugated to PE-Cy7 (Figure 1C). The expression of CD16/32 is not found on LECs but is found on LSECs. Fc block uses the CD16/32 antibody to minimize the Fc receptor binding and nonantigen specific binding of immunoglobulins to the Fc receptors. Since CD16/32 was used to visualize LSECs, the non-specific binding of immunoglobulins was higher and CD146 was optimized as an alternative to measure LSECs. Therefore, we tested CD146 conjugated to V450, a vascular endothelial cell marker expressed highly by LSECs and other vascular endothelial cells20 but with low to no expression by LECs23. We first optimized the staining of CD146 using FMO and isotype controls for both CD16/32 and CD146 (Figure 2A). If we evaluated only the CD16/32+ cells or the CD16/32- cells, the CD16/32+ cells were all CD146+ and PDPN- while the CD16/32- cells were primarily CD146- and either PDPN- or PDPN+ (Figure 2B). To confirm this staining, FMO was used. By removing PDPN from the stain, the PDPN+ population disappeared (Figure 2C). Interestingly, there were no CD146+ PDPN+ cells, confirming that CD146 is not or very lowly expressed by PDPN+ LECs. Thus, we are confident that either of these markers can be used to gate out vascular endothelial cells in the liver.
To further validate that PDPN is an appropriate marker for LECs, liver sections from mice were stained with both PDPN (green) and F4/80 (brown) (Figure 3A). Neither the vascular endothelium nor macrophages expressed PDPN in the murine liver. We were also able to distinguish cholangiocytes, which can stain positive for PDPN, from lymphatic vessels, based on the distinct nuclear structures of the bile ducts and by cytokeratin 7 staining (Figure 3B; red: cytokeratin 7, green: PDPN). Since multiple markers are used for flow cytometry, such as CD31, which is not expressed by cholangiocytes24, we are confident that we are removing these cells from the analysis, thereby confirming that cells from the liver that are CD45-, CD31+, CD146lo/neg, CD16/32-, and PDPN+ are LECs. Finally, to provide evidence that the stained cells are LECs, we sorted this population of cells and used qualitative real-time polymerase chain reaction to evaluate the Vegfr3 and prox-1 expressions. Both Vegfr3 and prox-1 were evaluated, as these transcripts are expressed by other cells in the liver&#8212;prox1 by hepatocytes and Vegfr3 by LSECS (among others) &#8212; but no other cells besides LECs express both. The expression of both these markers was significantly higher in the sorted population than in the cultured murine macrophage cell line (RAW264.7) which does not normally express these markers but is similar in expression to cultured murine LECs (Figure 3C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58621/58621fig1.jpg" /> 
Figure 1: Representative flow cytometry analysis of collagenase I-II-dispase and collagenase IV-digested murine liver tissue. (A) This panel shows the gating strategy, fluorescence minus one staining, and isotype controls for the procedure. (B) This panel shows the LYVE-1 staining of CD16/32- PDPN+ cells. (C) This panel shows the final flow cytometry gate after digesting mouse livers with either collagenase I-II-dispase (e.g., Liberase DL) or collagenase IV and evaluating CD16/32 PerCP X PDPN PE-Cy7 where LECs are CD16/32- and PDPN+. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58621/58621fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58621/58621fig2.jpg" /> 
Figure 2: Identification of CD146 and PDPN as appropriate markers for liver lymphatic endothelial cells. (A) This panel shows fluorescence minus one and isotype controls for CD16/32 (left) and CD146 (right). (B) This panel shows gated CD16/32 positive or negative cells determined from panel A. Shown is CD146XPDPN from both populations. (C) This panel shows fluorescence minus one for PDPN to demonstrate that the staining is absent when the antibody is not added. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58621/58621fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58621/58621fig3.jpg" /> 
Figure 3: Identification of flow PDPN+ cells as lymphatic endothelial cells. (A) This panel shows representative immunohistochemistry from a mouse liver stained with PDPN (blue/green) and F4/80 (brown). The lymphatic vessel (LV), blood vessel (BV), and macrophage (MF) are labeled. Scale bar = 100 &#181;m. Formalin-fixed paraffin-embedded tissue was deparaffinized for 20 min in xylene. Tissues were hydrated to water through a gradient of ethanol, and antigen retrieval was performed using pH 6 antigen retrieval buffer in a pressure cooker for 15 min. Tissue was blocked using 0.1% BSA and stained using anti-mouse PDPN and anti-mouse F4/80 for 1 h at room temperature. Anti-hamster IgG HRP and anti-rabbit IgG HRP were used as secondary antibodies. 3,3&#39;-Diaminobenzidine (DAB)+ and Vina Green were used to detect the F4/80 and PDPN, respectively. Tissue was counterstained with hematoxylin and imaged on a Microscope. (B) This panel shows the same experiment performed as in panel A, except here, PDPN is shown in green, cytokeratin 7 in red, and 4&#8242;,6-diamidino-2-phenylindole (DAPI) in blue. Scale bar = 100 &#181;m. Tissue was blocked using 5% donkey and 5% goat serum and stained using anti-mouse PDPN (8.1.1) 1:100 and anti-mouse cytokeratin 7 1:200 for 1 h at room temperature. Anti-hamster IgG AF647 and anti-rabbit IgG-PE were used as secondary antibodies. Tissue was counterstained with DAPI and imaged. Scale bar = 100 &#181;m. The lymphatic vessel (LV), blood vessel (BV), and bile duct (BD) are labeled. (C) This panel shows a log fold change in Vegfr3 and prox-1 expression from sorted liver LECs based on the staining in the protocol (CD45-, CD31+, CD146lo/neg, and PDPN+) compared to RAW cells or primary murine lymph node LECs. Sorted cells were passed through a biopolymer-shredding column, RNA was extracted using an RNA extraction kit, and cDNA was made using a reverse transcription kit. Transcript abundance was normalized to the housekeeping gene, Gapdh, for every sample. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58621/58621fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Digestion of the Murine Liver for a Flow Cytometric Analysis of Lymphatic Endothelial Cells at JoVE.com

Digestion of the Murine Liver for a Flow Cytometric Analysis of Lymphatic Endothelial Cells

The goal of this protocol is to identify lymphatic endothelial cell populations within the liver using described markers. We utilize collagenase IV and DNase and a gentle mincing of tissue, combined with flow cytometry, to identify a distinct population of lymphatic endothelial cells.

Within the liver, lymphatic vessels are found within the portal triad, and their described function is to remove interstitial fluid from the liver to the ...

digestion-murine-liver-for-flow-cytometric-analysis-lymphatic

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

10.6K Views.  University of Colorado Anschutz Medical Campus, School of Medicine. The goal of this protocol is to identify lymphatic endothelial cell populations within the liver using described markers. We utilize collagenase IV and DNase and a gentle mincing of tissue, combined with flow cytometry, to identify a distinct population of lymphatic endothelial cells.

Video: Digestion of the Murine Liver for a Flow Cytometric Analysis of Lymphatic Endothelial Cells

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 ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

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

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

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

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

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

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

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

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

Isolation and Flow Cytometric Characterization of Murine Small Intestinal Lymphocytes

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both microbial and dietary. Given an increasing understanding that these luminal antigens help shape the immune response and that education of immune cells within the intestine is critical for a number of systemic diseases, there has been increased interest in characterizing the intestinal immune system. However, many published protocols are arduous and time-consuming. We present here a simplified protocol for the isolation of lymphocytes from the small-intestinal lamina propria, intraepithelial layer, and Peyer&#39;s patches that is rapid, reproducible, and does not require laborious Percoll gradients. Although the protocol focuses on the small intestine, it is also suitable for analysis of the colon. Moreover, we highlight some aspects that may need additional optimization depending on the specific scientific question. This approach results in the isolation of large numbers of viable lymphocytes that can subsequently be used for flow cytometric analysis or alternate means of characterization.

There is growing interest in the quantitative characterization of intestinal lymphocytes owing to increasing recognition that these cells play a critical role in a variety of intestinal and systemic diseases. In this protocol, we describe how to isolate single cell populations from different small-intestinal compartments for subsequent flow cytometric characterization.

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both ...

Flow Cytometric Analysis of Particle-bound Bet v 1 Allergen in PM10

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in diameter. In addition to a characterization of forward and sideward scatter properties, it enables the use of fluorescent labeled markers like antibodies to detect respective structures. Using indirect antibody staining, flow cytometry is employed here to quantify birch pollen allergen (precisely Bet v 1)-loaded particles of 0.5 to 10 &#181;m in diameter in inhalable particulate matter (PM10, particle size &#8804;10 &#181;m in diameter). PM10 particles may act as carriers of adsorbed allergens possibly transporting them to the lower respiratory tract, where they could trigger allergic reactions.
So far the allergen content of PM10 has been studied by means of enzyme linked immunosorbent assays (ELISAs) and scanning electron microscopy. ELISA measures the dissolved and not the particle-bound allergen. Compared to scanning electron microscopy, which can visualize allergen-loaded particles, flow cytometry may additionally quantify them. As allergen content of ambient air can deviate from birch pollen count, allergic symptoms might perhaps correlate better with allergen exposure than with pollen count. In conjunction with clinical data, the presented method offers the opportunity to test in future experiments whether allergic reactions to birch pollen antigens are associated with the Bet v 1 allergen content of PM10 particles &#62;0.5 &#181;m.

Here, we present a protocol to quantify allergen-loaded particles by flow cytometry. Ambient particulate matter particles may act as carriers of adsorbed allergens. We show here that flow cytometry, a method widely used to characterize suspended solids &#62;0.5 &#181;m in diameter, can be used to measure these allergen-loaded particles.

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in ...

Flow Cytometric Analysis of Natural Killer Cell Lytic Activity in Human Whole Blood

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and reproducibility of this assay can be considered unstable, either because of user&#39;s errors or because of the sensitivity of NK cells to experimental manipulation. To eliminate these issues, a workflow that reduces them to a minimum was established and is presented here. To illustrate, we obtained blood samples, at various time points, from runners (n = 4) that were submitted to an intense bout of exercise. First, NK cells are simultaneously identified and isolated through CD56 tagging and magnetic-based sorting, directly from whole blood and from as little as one milliliter. The sorted NK cells are removed of any reagent or capping antibodies. They can be counted in order to establish an accurate NK cell count per milliliter of blood. Secondly, the sorted NK cells (effectors cells or E) can be mixed with 3,3&#39;-Diotadecyloxacarbocyanine Perchlorate (DiO) tagged K562 cells (target cells or T) at an assay-optimal 1:5 T:E ratio, and analyzed using an imaging flow-cytometer that allows for the visualization of each event and the elimination of any false positive or false negatives (such as doublets or effector cells). This workflow can be completed in about 4 h, and allows for very stable results even when working with human samples. When available, research teams can test several experimental interventions in human subjects, and compare measurements across several time points without compromising the data&#39;s integrity.

This work describes an advanced workflow for the accurate and fast determination of NK (Natural Killer) cell count and NK cell cytotoxicity in human blood samples.

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and ...

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 ...

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 ...