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>

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<li>Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Immunomodulatory+roles+of+lymphatic+vessels+in+cancer+progression%5BTitle%5D)+AND+%22Cancer+Immunology+Research%22%5BJournal%5D)">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/pubmed?term=((Cutting+edge%3A+lymphatic+vessels%2C+not+blood+vessels%2C+primarily+mediate+immune+rejections+after+transplantation%5BTitle%5D)+AND+%22Journal+of+Immunology%22%5BJournal%5D)">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/pubmed?term=((Migratory+Dendritic+Cells+acquire+archived+antigen+from+Lymphatic+Endothelial+Cells+for+antigen+presentation+during+lymph+node+contraction%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Migratory Dendritic Cells acquire archived antigen from Lymphatic Endothelial Cells for antigen presentation during lymph node contraction.</a> Nature Communications. 8, 2034(2017).</li>
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<li>Yokomori, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphatic+marker+podoplanin%2FD2-40+in+human+advanced+cirrhotic+liver--re-evaluations+of+microlymphatic+abnormalities%5BTitle%5D)+AND+%22BMC+Gastroenterology%22%5BJournal%5D)">Lymphatic marker podoplanin/D2-40 in human advanced cirrhotic liver--re-evaluations of microlymphatic abnormalities.</a> BMC Gastroenterology. 10, 131(2010).</li>
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</ol>

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&amp;amp;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>100 &amp;mu;m 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 &amp;micro;L pipete tip</td><td>USA scientific</td><td>1110-1700</td><td></td></tr><tr><td>10 &amp;micro;L pipete tip</td><td>USA scientific</td><td>1111-3700</td><td></td></tr><tr><td>seriological 10 mL pipete</td><td>greiner bio-one</td><td>607107</td><td></td></tr><tr><td>seriological 5 mL 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

10.7K 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

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 ...

Isolation of Human Lymphatic Endothelial Cells by Multi-parameter Fluorescence-activated Cell Sorting

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity but little is known about their biology. Isolating highly pure human lymphatic endothelial cells (LECs) from diseased and healthy tissue would facilitate studies of the lymphatic endothelium at genetic, molecular and cellular levels. It is anticipated that these investigations may reveal targets for new therapies that may change the clinical management of these conditions. A protocol describing the isolation of human foreskin LECs and lymphatic malformation lymphatic endothelial cells (LM LECs) is presented. To obtain a single cell suspension tissue was minced and enzymatically treated using dispase II and collagenase II. The resulting single cell suspension was then labelled with antibodies to cluster of differentiation (CD) markers CD34, CD31, Vascular Endothelial Growth Factor-3 (VEGFR-3) and PODOPLANIN. Stained viable cells were sorted on a fluorescently activated cell sorter (FACS) to separate the CD34LowCD31PosVEGFR-3PosPODOPLANINPos LM LEC population from other endothelial and non-endothelial cells. The sorted LM LECs were cultured and expanded on fibronectin-coated flasks for further experimental use.

The goal of this protocol is to isolate lymphatic endothelial cells lining human lymphatic malformation cyst-like vessels and foreskins using fluorescence-activated cell sorting (FACS). Subsequent cell culturing and expansion of these cells permits a new level of experimental sophistication for genetic, proteomic, functional and cell differentiation studies.

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity ...

Medicine

Identification and Analysis of Mouse Erythroid Progenitors using the CD71/TER119 Flow-cytometric Assay

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of red cell formation is regulated by the hormone erythropoietin (Epo), whose synthesis is triggered by tissue hypoxia. A threat to adequate tissue oxygenation results in a rapid increase in Epo, driving an increase in erythropoietic rate, a process known as the erythropoietic stress response. The resulting increase in the number of circulating red cells improves tissue oxygen delivery. An efficient erythropoietic stress response is therefore critical to the survival and recovery from physiological and pathological conditions such as high altitude, anemia, hemorrhage, chemotherapy or stem cell transplantation. 
The mouse is a key model for the study of erythropoiesis and its stress response. Mouse definitive (adult-type) erythropoiesis takes place in the fetal liver between embryonic days 12.5 and 15.5, in the neonatal spleen, and in adult spleen and bone marrow. Classical methods of identifying erythroid progenitors in tissue rely on the ability of these cells to give rise to red cell colonies when plated in Epo-containing semi-solid media. Their erythroid precursor progeny are identified based on morphological criteria. Neither of these classical methods allow access to large numbers of differentiation-stage-specific erythroid cells for molecular study. Here we present a flow-cytometric method of identifying and studying differentiation-stage-specific erythroid progenitors and precursors, directly in the context of freshly isolated mouse tissue. The assay relies on the cell-surface markers CD71, Ter119, and on the flow-cytometric 'forward-scatter' parameter, which is a function of cell size. The CD71/Ter119 assay can be used to study erythroid progenitors during their response to erythropoietic stress in vivo, for example, in anemic mice or mice housed in low oxygen conditions. It may also be used to study erythroid progenitors directly in the tissues of genetically modified adult mice or embryos, in order to assess the specific role of the modified molecular pathway in erythropoiesis.

A flow-cytometric method for identification and molecular analysis of differentiation-stage-specific murine erythroid progenitors and precursors, directly in freshly &#x2013;harvested mouse bone marrow, spleen or fetal liver. The assay relies on cell-surface markers CD71, Ter119, and cell size.

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of ...

Biology

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 ...

Digestion of Whole Mouse Eyes for Multi-Parameter Flow Cytometric Analysis of Mononuclear Phagocytes

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. Innate immune cells, specifically mononuclear phagocytes, express overlapping cell surface markers, which makes identifying these populations a challenge. Multi-parameter flow cytometry allows for the simultaneous, quantitative analysis of multiple cell surface markers in order to differentiate monocytes, macrophages, microglia, and dendritic cells in mouse eyes. This protocol describes the enucleation of whole mouse eyes, ocular dissection, digestion into a single cell suspension, and staining of the single cell suspension for myeloid cell markers. Additionally, we explain the proper methods for determining voltages using single color controls and for delineating positive gates using fluorescence minus one controls. The major limitation of multi-parameter flow cytometry is the absence of tissue architecture. This limitation can be overcome by multi-parameter flow cytometry of individual ocular compartments or complimentary immunofluorescence staining. However, immunofluorescence is limited by its lack of quantitative analysis and reduced number of fluorophores on most microscopes. We describe the use of multi-parametric flow cytometry to provide highly quantitative analysis of mononuclear phagocytes in laser-induced choroidal neovascularization. Additionally, multi-parameter flow cytometry can be used for the identification of macrophage subsets, fate mapping, and cell sorting for transcriptomic or proteomic studies.

This protocol provides a method to digest whole eyes into a single cell suspension for the purpose of multi-parameter flow cytometric analysis in order to identify specific ocular mononuclear phagocytic populations, including monocytes, microglia, macrophages, and dendritic cells.

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. ...

Flow Cytometric Analysis for Identification of the Innate and Adaptive Immune Cells of Murine Lung

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while suppressing unwanted reactions to environmental antigens. Lungs host several populations of innate and adaptive immune cells that provide immune surveillance but also mediate protective immune responses. These cells, which keep the healthy pulmonary immune system in balance, also participate in several pathological conditions such as asthma, infections, autoimmune diseases, and cancer. Selective expression of surface and intracellular proteins provides unique immunophenotypic properties to the immune cells of the lung. Consequently, flow cytometry has an instrumental role in the identification of such cell populations during steady-state and pathological conditions. This paper presents a protocol that describes a consistent and reproducible method to identify the immune cells that reside in the lungs of healthy mice under steady-state conditions. However, this protocol can also be used to identify changes in these cell populations in various disease models to help identify disease-specific changes in the lung immune landscape.

In this study, we present an effective and reproducible protocol to isolate the immune populations of the murine respiratory system. We also provide a method for the identification of all innate and adaptive immune cells that reside in the lungs of healthy mice, using a 9-color-based flow cytometry panel.

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while ...

Murine Model for Studying Postoperative Ileus through Intestinal Manipulation

Postoperative Ileus Murine Model

Most patients experience postoperative ileus (POI) after surgery, which is associated with increased morbidity, mortality, and hospitalization time. POI is a consequence of mechanical damage during surgery, resulting in disruption of motility in the gastrointestinal tract. The mechanisms of POI are related to aberrant neuronal sensitivity, impaired epithelial barrier function, and increased local inflammation. However, the details remain enigmatic. Therefore, experimental murine models are crucial for elucidating the pathophysiology and mechanism of POI injury and for the development of novel therapies.
Here, we introduce a murine model of POI generated via intestinal manipulation (IM) that is similar to clinical surgery; this is achieved by mechanical damage to the small intestine by massaging the abdomen 1-3 times with a cotton swab. IM delayed gastrointestinal transit 24 h after surgery, as assessed by FITC-dextran gavage and fluorescence detection of the segmental digestive tract. Moreover, tissue swelling of the submucosa and immune cell infiltration were investigated by hematoxylin and eosin staining and flow cytometry. Proper pressure of the IM and a hyperemic effect on the intestine are critical for the procedure. This murine model of POI can be utilized to study the mechanisms of intestinal damage and recovery after abdominal surgery.

We describe a murine model of postoperative ileus generated via intestinal manipulation. Gastrointestinal transit function, pathologic changes, and immune cell activation were assessed 24 h after surgery.

Most patients experience postoperative ileus (POI) after surgery, which is associated with increased morbidity, mortality, and hospitalization time. POI ...

Isolation of Immune and Non&mdash;Immune Cells from Mouse Lungs for Flow Cytometry

Isolation of Live Myeloid and Epithelial Cell Populations from the Mouse Lung

The lung is continuously exposed to pathogens and other noxious environmental stimuli, rendering it vulnerable to damage, dysfunction, and the development of disease. Studies utilizing mouse models of respiratory infection, allergy, fibrosis, and cancer have been critical to reveal mechanisms of disease progression and identify therapeutic targets. However, most studies focused on the mouse lung prioritize the isolation of either immune cells or epithelial cells, rather than both populations concurrently. Here, we describe a method for preparing a comprehensive single-cell suspension of both immune and non-immune populations suitable for flow cytometry and fluorescence-activated cell sorting. These populations include epithelial cells, endothelial cells, fibroblasts, and a variety of myeloid cell subsets. This protocol entails bronchoalveolar lavage and subsequent inflation of the lungs with dispase. Lungs are then digested in a liberase mixture. This method of processing liberates a variety of diverse cell types and results in a single-cell suspension that does not require manual dissociation against a filter, promoting cell survival and yielding high numbers of live cells for downstream analyses. In this protocol, we also define gating schemes for epithelial and myeloid cell subsets in both na&#239;ve and influenza-infected lungs. Simultaneous isolation of live immune and non-immune cells is key for interrogating intercellular crosstalk and gaining a deeper understanding of lung biology in health and disease.

This protocol details the isolation of live immune and non-immune populations from the mouse lung at a steady state and following influenza infection. It also provides gating strategies for identifying epithelial and myeloid cell subsets.

The lung is continuously exposed to pathogens and other noxious environmental stimuli, rendering it vulnerable to damage, dysfunction, and the development ...

Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. LSECs have a distinct morphology characterized by the presence of fenestrae and the absence of basement membrane. LSECs play essential roles in many pathological disorders in the liver, including metabolic dysregulation, inflammation, fibrosis, angiogenesis, and carcinogenesis. However, little has been published about the isolation and characterization of the LSECs. Here, this protocol discusses the isolation of LSEC from both healthy and nonalcoholic fatty liver disease (NAFLD) mice. The protocol is based on collagenase perfusion of the mouse liver and magnetic beads positive selection of nonparenchymal cells to purify LSECs. This study characterizes LSECs using specific markers by flow cytometry and identifies the characteristic phenotypic features by scanning electron microscopy. LSECs isolated following this protocol can be used for functional studies, including adhesion and permeability assays, as well as downstream studies for a particular pathway of interest. In addition, these LSECs can be pooled or used individually, allowing multi-omics data generation including RNA-seq bulk or single cell, proteomic or phospho-proteomics, and Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq), among others. This protocol will be useful for investigators studying LSECs&#39; communication with other liver cells in health and disease and allow an in-depth understanding of the role of LSECs in the pathogenic mechanisms of acute and chronic liver injury.

Here we outline and demonstrate a protocol for primary mouse liver sinusoidal endothelial cell (LSEC) isolation. The protocol is based on liver collagenase perfusion, nonparenchymal cell purification by low-speed centrifugation, and CD146 magnetic bead selection. We also phenotype and characterize these isolated LSECs using flow cytometry and scanning electron microscopy.

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. ...

Isolating Endothelial Cells from Murine Dermal Lymphatic Capillaries

Murine Dermal Lymphatic Endothelial Cell Isolation

The lymphatic system participates in the regulation of immune surveillance, lipid absorption, and tissue fluid balance. The isolation of murine lymphatic endothelial cells is an important process for lymphatic research, as it allows the performance of in vitro and biochemical experiments on the isolated cells. Moreover, the development of Cre-lox technology has enabled the tissue-specific deficiency of genes that cannot be globally targeted, leading to the precise determination of their role in the studied tissues. The dissection of the role of certain genes in lymphatic physiology and pathophysiology requires the use of lymphatic-specific promoters, and thus, the experimental verification of the expression levels of the targeted genes.
Methods for efficient isolation of lymphatic endothelial cells from wild-type or transgenic mice enable the use of ex vivo and in vitro assays to study the mechanisms regulating the lymphatic functions and the identification of the expression levels of the studied proteins. We have developed, standardized and present a protocol for the efficient isolation of murine dermal lymphatic endothelial cells (DLECs) via magnetic bead purification based on LYVE-1 expression. The protocol outlined aims to equip researchers with a tool to further understand and elucidate important players of lymphatic endothelial cell functions, especially in facilities where fluorescence-activated cell sorting equipment is not available.

Author Spotlight: Magnetic Bead-Based Isolation of Murine Dermal Lymphatic Endothelial Cells

This paper describes a method for magnetic bead-based isolation of murine endothelial cells from dermal lymphatic capillaries. The isolated lymphatic endothelial cells can be used for downstream in vitro experiments and protein expression analysis.

The lymphatic system participates in the regulation of immune surveillance, lipid absorption, and tissue fluid balance. The isolation of murine lymphatic ...