This work was supported by the Hellenic Foundation for Research and Innovation (00376), the National Institutes of Health, the National Cancer Institute (NCI) [Grant R15CA231339], the Texas Tech University Health Sciences Center (TTUHSC) School of Pharmacy Office of the Sciences grant (to C.M.M.), and by the College of Pharmacy, University of Louisiana Monroe start-up funding, the National Institutes of Health (NIH) through the National Institute of General Medical Science [Grant P20 GM103424-21] and the Research Competitiveness Subprogram (RCS) of the Louisiana Board of Regents through the Board of Regents Support Fund (LEQSF(2021-24)-RD-A-23) (to G.M.). The common TTUHSC equipment used was obtained through the Cancer Prevention Research Institute of Texas (CPRIT) Grants RP190524 and RP200572. The funders had no role in the study design, the decision to write, or preparation of the manuscript. The graphical abstract in Figure 1A was created with BioRender.com.

Animal studies were performed according to the approved protocols by the Institutional Animal Care and Use Committee (IACUC) of Texas Tech University Health Sciences Center (TTUHSC) for the experiments at TTUHSC and by the Veterinary Administration of the Prefecture of Western Greece according to Directive 2010/63 for the experiments at the University of Patras. Diligently follow waste disposal regulations when disposing of animal waste materials.
1. Isolation of mouse dermis
<ol>
	<li>Day 1 
	NOTE: On Day 1; get the following materials and equipment ready: anesthesia chamber, animal hair clipper, isoflurane, fine forceps, 0.2 &#181;m syringe filters.
	<ol>
		<li>Prepare a 2 mg/mL Dispase solution in DMEM with 1x Antibiotic-Antimycotic Solution and filter-sterilize it. 
		NOTE: The use of medium or PBS can be replaced by HBSS, as it has been successfully used in other reported&#160;protocols12. The 1x Antibiotic-Antimycotic Solution can be substituted by the addition of penicillin (100 I.U./mL), streptomycin (100 &#181;g/mL), and amphotericin (2.5 &#181;g/mL).</li>
		<li>Anesthetize the mouse by placing the mouse in an anesthesia chamber with isoflurane administration (0.5-2% isoflurane, flow: 1 L/min). After the mouse stops moving, transfer it out of the cage, and continue anesthesia adjusting a nose cone with a similar isoflurane flow. Toe-pinch the animal to confirm sufficient anesthesia levels and constantly monitor the mouse to ensure deep anesthesia is preserved (no motility). Clip approximately 9 cm2 of the dorsal and ventral fur (mouse back and abdomen respectively) longitudinally along the midline, using a hair clipper. Let the mouse recover from anesthesia and return it to the cage. 
		NOTE: Hair removal (depilation) cream can be used instead. Make sure the fur is completely removed in the skin areas to be isolated while maintaining the integrity of the skin. Try to clip as extended&#160;skin area as possible. To ensure that potential inflammation from the hair removal is resolved, cell isolation must take place the following day. To increase the number of isolated cells pool 2 to 4 mice per group.</li>
	</ol>
	</li>
	<li>Day 2 
	NOTE: Keep the following materials and equipment ready: euthanasia chamber, biosafety cabinet, CO2 incubator, ethanol, fine scissors, fine forceps, blunt forceps, 100 mm tissue culture plate, Dispase, DMEM, 1x Antibiotic-Antimycotic Solution.
	<ol>
		<li>Euthanize the mouse by CO2 administration and place it in a supine position, exposing the clipped area. Spray 70% ethanol as a disinfectant. 
		NOTE: From this step onwards, all steps must be performed under sterile conditions (in a biosafety cabinet).</li>
		<li>Place a sterile 100 mm Tissue Culture Plate on ice.</li>
		<li>Make a 0.5 inch superficial cut at the abdomen and cut longitudinally along the midline to the neck and the lower abdomen. Make lateral cuts to be able to flap open the skin. 
		NOTE: The incision size and the skin area isolated depend on the mouse size.</li>
		<li>Using blunt forceps, carefully separate the skin from the surrounding tissues. Do this going around the entire torso of the mouse, cutting the skin laterally around the neck and the lower abdomen.</li>
		<li>Carefully place the skin into the 100 mm Tissue Culture Plate, with the dermis side facing down, keeping the skin as stretched as possible. Cover the plates and keep them on ice. Complete the isolation of all mice from the group.</li>
		<li>After the skin is extracted from all mice, incubate plates for 10 min at room temperature to flatten the explant.</li>
		<li>Add 6 mL of Dispase solution to the plate, making sure the skin is completely immersed and incubate overnight with mild agitation at 4 &#176;C. 
		NOTE: Instead of Dispase, trypsin digestion can also be performed12, followed by Liberase instead of Collagenase Type II treatment (below).</li>
	</ol>
	</li>
</ol>
2. Cell isolation from the dermis
<ol>
	<li>Day 3 
	NOTE: Keep the following materials and equipment ready: centrifuge, CO2 incubator, biosafety cabinet, scissors, DMEM, 1x Antibiotic-Antimycotic Solution, Collagenase Type II, PBS, 50 mL conical tube, 100 mm and 60 mm tissue culture plates, DNAse I solution, Collagen Type I, 70 &#181;m and 40 &#181;m cell strainers, 0.2 &#181;m syringe filters, LEC medium.
	<ol>
		<li>Prepare 500 mL of 1x PBS (1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4, pH 7.4) and autoclave for 20 min at 15 psi or filter-sterilize. Alternatively, use commercially available sterile PBS.</li>
		<li>Prepare 10 mL of 2 mg/mL Collagenase Type II solution in DMEM with 1x Antibiotic-Antimycotic Solution and filter-sterilize it. 
		NOTE: Perform the next steps under a sterile hood to decrease the chances of sample contamination.</li>
		<li>Use a pipette to remove the liquid from the dish.</li>
		<li>Use forceps to separate the dermis (bottom of the skin) from the epidermis (top) and remove any obvious adipose tissue. Transfer the dermis to a clean Tissue Culture plate.</li>
		<li>Add 1 mL of DMEM with 1x Antibiotic-Antimycotic Solution in the clean Tissue Culture Plate and tilt it so that the liquid accumulates on one side of the plate.</li>
		<li>Mince the dermis into 1-2 mm2 pieces and transfer them to a new 50 mL conical tube. 
		NOTE: This step must ideally be completed within 4-5 min, as longer processing is not expected to provide higher yield or cell survival.</li>
		<li>Add 10 mL of the Collagenase Type II solution.</li>
		<li>Add 50 &#181;L (25 units/mL of the solution) of DNAse I solution (reduces cell clumping) and digest the dermis solution for 2 h at 37 &#176;C with continuous agitation (use a rotating/shaking incubator at 30-50 rpm). 
		NOTE: Check periodically to identify whether most of the dermis has been digested. If not, increase the digestion time (1 extra h maximum) and agitation speed, if possible.</li>
		<li>Coat 60 mm Tissue Culture Plates with 2 mL of 10 &#181;g/mL Collagen Type I in 0.02 M Acetic Acid for &#62;20 min at 37 &#176;C. Wash 2x with sterile PBS.</li>
		<li>Filter the cell suspension using a 70 &#181;m cell strainer and centrifuge at 250 &#215; g for 5 min. Discard the supernatant.</li>
		<li>Resuspend in 10 mL of DMEM with 1x Antibiotic-Antimycotic Solution.</li>
		<li>Filter the cell suspension using a 40 &#181;m cell strainer and centrifuge at 250 &#215; g for 5 min. Discard the supernatant.</li>
		<li>Resuspend in 1 mL of complete LEC media (Endothelial Cell Growth Base Medium with Supplement).</li>
		<li>Plate the cells in the collagen-coated 60 mm Tissue Culture Plate in complete LEC media. This will be Day 0 of cell culture.</li>
		<li>Replace the media after every 2 days by washing cells with 1x PBS twice and adding fresh LEC media.</li>
		<li>Repeat step 2.1.15 until the cells are 80-90% confluent. 
		&#8203;NOTE: If the purification is performed before the cells are 80-90% confluent, fewer lymphatic endothelial cells will be isolated. This process takes about 7 to 10 days.</li>
	</ol>
	</li>
</ol>
3. Magnetic Bead Coating with LYVE-1 antibody
NOTE: One day before the purification step, get the following materials and equipment ready: 4 &#176;C Refrigerator, Rotor, magnetic separator, PBS, Anti-Rabbit IgG Magnetic Beads, microcentrifuge tubes, 0.2 &#181;m syringe filters, Magnetic Bead Coating Solution, LYVE-1 antibody.
<ol>
	<li>Prepare the Magnetic Bead Coating Solution: 20 mL of PBS with 0.1% BSA and 2 mM EDTA and filter-sterilize it. Store it at 4 &#176;C. 
	NOTE: More than a few days of storage is not recommended; check for precipitates or contamination before use.</li>
	<li>Pipette 500 &#181;L of Anti-Rabbit IgG Magnetic Beads to a microcentrifuge tube and add 1 mL of ice-cold Magnetic Bead Coating Solution.</li>
	<li>Use a magnetic separator to wash the beads.</li>
	<li>Repeat the washes 2x with 1 mL of the ice-cold Magnetic Bead Coating Solution.</li>
	<li>Resuspend the beads in 500 &#181;L of Magnetic Bead Coating Solution and make aliquots of 150 &#181;L in microcentrifuge tubes.</li>
	<li>Add 15 &#181;g of the LYVE-1 antibody to each tube (15 &#181;L from 1 mg/mL antibody stock concentration).</li>
	<li>Incubate overnight at 4 &#176;C&#160;with continuous agitation.</li>
</ol>
4. Lymphatic endothelial cell purification
NOTE: Get the following materials and equipment ready: CO2 incubator, shaker, timer, parafilm, DMEM, 1x Antibiotic-Antimycotic Solution, BSA, PBS, Magnetic Bead Coating Solution, trypsin-EDTA, 100 mm or 60 mm tissue culture plates, 0.2 &#181;m syringe filters, Collagen Type I, Acetic acid, LEC medium.
<ol>
	<li>Prepare 50 mL of DMEM with 1x Antibiotic-Antimycotic Solution and 0.1% BSA and filter-sterilize it.</li>
	<li>Coat 60 mm Tissue Culture Plates with 10 &#181;g/mL Collagen Type I in 0.02 M Acetic Acid for &#62;20 min at 37 &#176;C. Wash 2x with sterile PBS.</li>
	<li>Wash the precoated beads with 1 mL of Magnetic Bead Coating Solution, using a magnetic separator.</li>
	<li>Resuspend the beads in 150 &#181;L of DMEM with 0.1% BSA. 
	NOTE: The beads can now be stored at 4 &#176;C for up to 1 month.</li>
	<li>Wash the cells 2x with PBS.</li>
	<li>Add 3 mL of DMEM with 0.1% BSA (for a 60 mm Tissue Culture Plate) and 50 &#181;L of the antibody-conjugated beads to the cells. 
	NOTE: If the cells from two or more mice are pooled, use a 100 mm Tissue Culture Plate and add 6 mL of DMEM with 0.1% BSA.</li>
	<li>Seal the dish with parafilm.</li>
	<li>Place the dish on a shaker agitating at 10 RPM at RT and incubate for exactly 10 min. 
	NOTE: Longer incubation will result in non-specific binding. To ensure that the beads interact with the entire plate surface, manually rotate the plate after every few minutes as it continues to shake.</li>
	<li>Discard the medium and wash the cells once with PBS. Quickly inspect the cells to verify the presence of LEC colonies on the plate.</li>
	<li>Trypsinize the cells: use 1 mL of trypsin-EDTA and incubate for 3 min at 37 &#176;C. Inspect the cells for successful cell detachment.</li>
	<li>Neutralize the solution with 2 mL of complete LEC media and transfer the solution to a 5 mL sterile polypropylene tube.</li>
	<li>Using a magnetic separator wash cells 4 x 3 mL of DMEM with 0.1% BSA to remove unbound cells.</li>
	<li>Resuspend the remaining bead-bound cells in complete LEC media and plate onto the previously Collagen-coated 60 mm Tissue Culture Plates.</li>
	<li>Incubate the cells at 37 &#176;C and change the media every other day after two PBS washes each time.</li>
	<li>Culture the cells till they reach confluency (it normally takes 7 to 10 days).</li>
	<li>At that point, check cells under the microscope for confluency and proceed with preparing cells for flow cytometry. If a high F4/80+ percentage is identified, then repeat the purification step with LYVE-1 (Step 4: Lymphatic Endothelial Cell Purification) again or ideally another lymphatic marker (i.e., podoplanin14). 
	&#8203;NOTE: The high expression of the protein targeted by the antibody on the magnetic bead-purified cell population has been demonstrated15,16. However, for LYVE-1, it is important to perform a flow cytometry control for the identification of the percentage of other LYVE-1+ cells, mostly macrophages. LYVE-1 is only expressed in subsets of macrophages that are associated with tumors and wounds, whereas in healthy skin, LYVE-1 expression is more restricted to lymphatic endothelial cells17. Thus, the F4/80 screening is not mandatory, but rather dependent on the experimental context. The mice used for demonstration did not carry tumors or wounds, just potential inflammation from hair removal, but we included the F4/80 staining as a baseline of the percentage of LYVE-1+ macrophages.</li>
</ol>
5. Flow cytometry
NOTE: Get the following materials and equipment ready: centrifuge, CO2 incubator, timer, hemocytometer, EDTA solution, 1x Antibiotic-Antimycotic Solution, BSA, PBS, trypsin-EDTA, polypropylene tubes, 0.2 &#181;m syringe filters, 70% ethanol, F4/80 FITC-conjugated antibody.
<ol>
	<li>Prepare PBS containing 0.1% BSA and filter-sterilize it.</li>
	<li>Prepare 1 mM EDTA solution in PBS.</li>
	<li>Trypsinize the cells: incubate the cells with 1 mL of 1 mM EDTA solution for 5 min at 37 &#176;C. 
	NOTE: Incubation with EDTA solution is a milder method of cell detachment, which preserves better the transmembrane proteins and is preferred, but it requires longer incubation periods. Alternatively, trypsin-EDTA solution can be used for 1 min incubation at 37 &#176;C, but this can affect antibody affinity for flow cytometry applications.</li>
	<li>Neutralize the solution with 2 mL of complete LEC media and transfer the solution to a 5 mL sterile polypropylene tube.</li>
	<li>Centrifuge the samples at 250 &#215; g for 5 min.</li>
	<li>Wash with PBS.</li>
	<li>Repeat the centrifugation and PBS wash.</li>
	<li>Resuspend the cells in PBS containing 0.1% BSA and count them using a hemocytometer or alternative method. Aliquot ~1 &#215; 106 cells into polypropylene tubes with a final volume of 100 &#181;l. From this step onwards, perform all the steps at 4 &#176;C. 
	NOTE: The cells can be fixed before staining or stained while alive. Fixation can be suboptimal for flow cytometry, as some antibodies may not detect the antigens after fixation; therefore, we recommend running flow cytometry immediately after the previous step and skipping the fixation step. If that is not possible, fixation prior to the flow cytometry experiment will allow a delay in the flow cytometry step. If fixation is to be performed, the performance of the antibodies in the context of fixed cells should be evaluated in advance.</li>
	<li>Perform fixation (optional step; if the cells are not to be fixed, proceed with immunostaining): resuspend the cells in ice-cold 70% ethanol and gently mix for 10 min on ice. 
	NOTE: Ethanol fixation can permeabilize the cells and lead to the release of GFP; however, paraformaldehyde-based fixation can destroy dyes used for flow cytometry and ideally can be avoided.</li>
	<li>Incubate for 15 min at room temperature.</li>
	<li>Wash cells 2x by centrifugation at 250 &#215; g for 5 min in 1x PBS.</li>
	<li>Proceed to the next step or resuspend the cells in 0.5-1 mL of 1x PBS and store at 4 &#176;C.</li>
</ol>
6. Immunostaining
NOTE: Handle the cells gently and avoid extensive pipetting, when possible, to maintain the cell membrane integrity and minimize cell death, ensuring that the cells are not forming clumps. From the isolated endothelial cells, prepare the following groups: unstained cells and stained cells with another endothelial cell marker (i.e., VEGFR3, podoplanin), F4/80, and both. Include commercially available mouse lymphatic endothelial cells and macrophages, if available, as positive controls. For demonstration purposes, and due to the endogenous tdTomato fluorescence (PE+), we have used FITC-conjugated F4/80 staining.
<ol>
	<li>Dilute the primary antibody in PBS with 0.1% BSA according to the recommended or optimized dilution. Add 0.5 &#181;g (1 &#181;L of antibody solution) of the F4/80 FITC-conjugated antibody to the 100 &#181;Lof the cell suspension.</li>
	<li>Resuspend the cells after the addition of the diluted fluorescent-conjugated antibody. Mix by flicking the polypropylene tube to ensure that there are no clumps and all cells are properly coated.</li>
	<li>Incubate for 1 h at room temperature (fixed cells) or 30 min on ice (live cells) in the dark.</li>
	<li>Wash cells 2x by centrifuging them at 250 &#215; g for 5 min at 4 &#176;C (for live cells) or at room temperature (for fixed cells), using PBS with 0.1% BSA in the dark.</li>
	<li>For non-fluorescently-conjugated antibodies
	<ol>
		<li>Dilute the fluorochrome-conjugated secondary antibody in the incubation buffer. For Alexa Flour antibodies, use a 1:500 dilution.</li>
		<li>Add the secondary antibody to 100 &#181;L of the cell suspension solution.</li>
		<li>Incubate for 30 min at room temperature (fixed cells) or on ice (live cells).</li>
		<li>Wash twice by centrifugation at 250 &#215; g for 5 min at 4 &#176;C (for live cells) or at room temperature (for fixed cells) in the incubation buffer.</li>
	</ol>
	</li>
	<li>Resuspend cells in 0.5 mL of 1x PBS and analyze them using a flow cytometer.</li>
	<li>Follow these gating strategy steps to identify the macrophage population within the LYVE-1-expressing cells:
	<ol>
		<li>Consider forward and side scattering to distinguish the cellular population.</li>
		<li>Consider forward scattering for cell size area and height to distinguish the single cells (Singlets).</li>
		<li>Take into account F4/80 (FITC) expression and RFP (PE) expression to identify macrophages (F4/80+) within the tdTomato (PE)-expressing isolated murine cells (due to the presence of a fluorescent tdTomato reporter in the specific mice, described below).</li>
	</ol>
	</li>
</ol>

Isolating Endothelial Cells from Murine Dermal Lymphatic Capillaries

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H., 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+dysfunction,+not+aplasia,+underlies+Milroy+disease.">Lymphatic dysfunction, not aplasia, underlies Milroy disease.</a> Microcirculation. 17 (4), 281-296 (2010).</li><li>Connell, F., Brice, G., Mortimer, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotypic+characterization+of+primary+lymphedema.">Phenotypic characterization of primary lymphedema.</a> Annals of the New York Academy of Sciences. 1131, 140-146 (2008).</li><li>Croteau, S. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kaposiform+lymphangiomatosis:+a+distinct+aggressive+lymphatic+anomaly.">Kaposiform lymphangiomatosis: a distinct aggressive lymphatic anomaly.</a> Journal of Pediatris. 164 (2), 383-388 (2014).</li><li>Ji, Y., Chen, S., Peng, S., Xia, C., Li, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kaposiform+lymphangiomatosis+and+kaposiform+hemangioendothelioma:+similarities+and+differences.">Kaposiform lymphangiomatosis and kaposiform hemangioendothelioma: similarities and differences.</a> Orphanet Journal of Rare Diseases. 14 (1), 165 (2019).</li><li>Kriehuber, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+characterization+of+dermal+lymphatic+and+blood+endothelial+cells+reveal+stable+and+functionally+specialized+cell+lineages.">Isolation and characterization of dermal lymphatic and blood endothelial cells reveal stable and functionally specialized cell lineages.</a> Journal of Experimental Medicine. 194 (6), 797-808 (2001).</li><li>Podgrabinska, S., 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=Molecular+characterization+of+lymphatic+endothelial+cells.">Molecular characterization of lymphatic endothelial cells.</a> Proceedings of the National Academy of Sciences of the United States of America. 99 (25), 16069-16074 (2002).</li><li>Garrafa, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation,+purification,+and+heterogeneity+of+human+lymphatic+endothelial+cells+from+different+tissues.">Isolation, purification, and heterogeneity of human lymphatic endothelial cells from different tissues.</a> Lymphology. 38 (4), 159-166 (2005).</li><li>Kazenwadel, J., Secker, G. A., Betterman, K. L., Harvey, N. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+assays+using+primary+embryonic+mouse+lymphatic+endothelial+cells+uncover+key+roles+for+FGFR1+signalling+in+lymphangiogenesis.">In vitro assays using primary embryonic mouse lymphatic endothelial cells uncover key roles for FGFR1 signalling in lymphangiogenesis.</a> PLoS One. 7 (7), e40497 (2012).</li><li>Steele, M. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T+cell+egress+via+lymphatic+vessels+is+tuned+by+antigen+encounter+and+limits+tumor+control.">T cell egress via lymphatic vessels is tuned by antigen encounter and limits tumor control.</a> Nature Immunology. 24 (4), 664-675 (2023).</li><li>Mammoto, 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=Effects+of+age-dependent+changes+in+cell+size+on+endothelial+cell+proliferation+and+senescence+through+YAP1.">Effects of age-dependent changes in cell size on endothelial cell proliferation and senescence through YAP1.</a> Aging. 11 (17), 7051-7069 (2019).</li><li>Regina, C., 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=Vascular+ageing+and+endothelial+cell+senescence:+Molecular+mechanisms+of+physiology+and+diseases.">Vascular ageing and endothelial cell senescence: Molecular mechanisms of physiology and diseases.</a> Mechanisms of Ageing and Development. 159, 14-21 (2016).</li><li>Muhleder, S., Fernandez-Chacon, M., Garcia-Gonzalez, I., Benedito, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endothelial+sprouting,+proliferation,+or+senescence:+tipping+the+balance+from+physiology+to+pathology.">Endothelial sprouting, proliferation, or senescence: tipping the balance from physiology to pathology.</a> Cellular and Molecular Life Sciences: CMLS. 78 (4), 1329-1354 (2021).</li></ol>

The authors declare no competing financial interests.

The lymphatic system is an important regulator of the homeostatic function of the body, with the most important functions being the maintenance of fluid plasma, removal of cellular metabolism byproducts and toxic molecules, lipid absorption, and immune cell trafficking1,19. The identification of appropriate markers has provoked a burst in new data in the lymphatics field, revealing novel functions of the lymphatic vasculature, such as their role in the repair and...

The lymphatic system plays a pivotal role in human physiology. It is considered a vital homeostatic factor, that facilitates important functions, such as the maintenance of tissue-plasma fluid balance, immune surveillance, and lipid absorption1, as well as recently-identified functions, such as the repair ability of the heart2 and regenerative capacity of bone and hematopoiesis3. Despite the significant role of the lymphatic system, several aspects of the role of the lymphatics, as well as the molecular mechanisms governing certain physiological parameters and responses remain elusive. Moreover, lymphatic vascular defects are triggering or deteriorating factors in certain pathophysiological conditions. Well-known examples are the primary and secondary lymphedemas, depending on the genetic or non-genetic origin of the disease, respectively, while the role of the lymphatic system on primary tumor growth and metastatic dissemination is pivotal, as it can act as a conduit for metastasis and modulator of immune functions1.
The lag in the study and knowledge of the lymphatic compared to the blood vasculature is mainly due to the later discovery of the lymphatic system in comparison with the blood vascular system and the delay in the identification of lymphatic-specific molecular markers. This has greatly improved in recent decades, leading to the alteration of the conventional picture of the lymphatics at steady state and in diseased conditions1. Similar to angiogenesis, lymphangiogenesis is the formation of new lymphatic vessels from pre-existing ones and plays a pivotal role in the development of a wide spectrum of diseases4,5,6. However, unlike angiogenesis, the investigation of lymphangiogenesis has been limited to mostly in vivo models focusing on developmental vessel formation and structural defects in pathological models. The isolation of the lymphatic endothelial cells is important for in vitro studies, and this can be provided by the presented protocol.
Several protocols for lymphatic endothelial isolation from mice have been developed, which require the use of fluorescence-activated cell sorting7. The advantage of the cell sorter, where available, is the higher degree of purity, contrary to bead-based isolation, with the latter providing a higher yield. The protocol utilizes the expression of lymphatic endothelial hyaluronan receptor 1 (LYVE-1), a lymphatic endothelial marker, important for cell trafficking within the lymphatic vessels4,8. Since LYVE-1 expression is limited to the lymphatic capillaries and not the collecting lymphatic vessels, the technique is suitable for the isolation of lymphatic endothelial cells specifically from the lymphatic capillaries, contrary to other lymphatic markers, such as podoplanin, which are expressed uniformly in all lymphatic endothelial cells9. For the protocol, other important lymphatic markers that were not transmembrane, such as Prox1, were excluded.
As the physiological outcome was lymphangiogenesis, which is usually initiated at the lymphatic capillaries, LYVE-1 was selected as a target due to its selectivity in these cells and because the selected antibody provided a high yield. The enzymes used were collagenase type II, which is generally known to be more effective in collagen dissociation than type I10, and dispase, a broader protease, regularly used for dermis-epidermis separation11; however, other enzymes for tissue separation have been reported12,13 and can be used. The goal of this protocol is to describe a method for dermal lymphatic endothelial cell isolation from lymphatic capillaries of adult mice with high yield using magnetic bead purification, especially in places where fluorescence-activated cell sorting is not readily available. The method is suitable for applications where the purity of lymphatic endothelial cells can be compromised for downstream applications.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 &#956;m Syringe Filters</td>
			<td>Fisher Scientific</td>
			<td>09-719C</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mm Tissue Culture Dishes</td>
			<td>Fisher Scientific</td>
			<td>FB012924</td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm Tissue Culture Dishes</td>
			<td>Fisher Scientific</td>
			<td>FB012921</td>
			<td></td>
		</tr>
		<tr>
			<td>Animal Hair Clipper</td>
			<td>Wahl</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic Solution 100x</td>
			<td>Fisher Scientific</td>
			<td>15240-062</td>
			<td></td>
		</tr>
		<tr>
			<td>Blunt Forceps</td>
			<td>Fine Science Tools</td>
			<td>11992-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A4919</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Strainer 40 &#956;m</td>
			<td>Fisher Scientific</td>
			<td>542040</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Strainer 70 &#956;m</td>
			<td>Fisher Scientific</td>
			<td>542070</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 Incubator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Collagen I, High Concentration Rat Tail</td>
			<td>Corning</td>
			<td>354249</td>
			<td>dilute in 0.02 M Acetic Acid in H2O</td>
		</tr>
		<tr>
			<td>Collagenase Type II</td>
			<td>Life Technologies</td>
			<td>17101-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Dispase</td>
			<td>Life Technologies</td>
			<td>17105-041</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Fisher Scientific</td>
			<td>11-995-073</td>
			<td></td>
		</tr>
		<tr>
			<td>DNAse I Solution (2,500 U/mL)</td>
			<td>Thermo Scientific</td>
			<td>90083</td>
			<td></td>
		</tr>
		<tr>
			<td>Dynal MPC-L Magnetic Particle Concentrator</td>
			<td>Invitrogen</td>
			<td>120-21D</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>3690</td>
			<td></td>
		</tr>
		<tr>
			<td>Endothelial Cell Growth Base Medium &#38; Supplement (LEC medium)</td>
			<td>R&#38;D Systems</td>
			<td>CCM027</td>
			<td></td>
		</tr>
		<tr>
			<td>Euthanasia chamber</td>
			<td>Euthanex Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11255-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors-Sharp</td>
			<td>Fine Science Tools</td>
			<td>14060-10</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti-mouse F4/80 Antibody</td>
			<td>Biolegend</td>
			<td>123107</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat Anti-Rabbit IgG Magnetic Beads</td>
			<td>New England Biolabs</td>
			<td>S1432S</td>
			<td></td>
		</tr>
		<tr>
			<td>LARC-A E-Z Anesthesia Induction Chamber</td>
			<td>Euthanex Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MagnaBind Goat Anti-Rabbit IgG Beads</td>
			<td>Thermo Scientific</td>
			<td>21356</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde Solution 4% in PBS</td>
			<td>Fisher Scientific</td>
			<td>AAJ19943K2</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Fisher Scientific</td>
			<td>SH30256FS</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit Anti-Mouse LYVE-1</td>
			<td>ReliaTech GmbH</td>
			<td>103-PA50</td>
			<td></td>
		</tr>
		<tr>
			<td>Rotating/Shaking Incubator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Round-Bottom Polypropylene Tubes</td>
			<td>Corning</td>
			<td>352063</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Filters w 0.2 &#956;m Pores</td>
			<td>Fisher Scientific</td>
			<td>09-719C</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Fisher Scientific</td>
			<td>25-300-120</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The method was developed to identify the protein expression of a small GTPase, RhoA, the coding gene of which was floxed in both alleles and deleted under the effect of the tamoxifen-inducible lymphatic endothelial-specific promoter Prox1-CreERT2 18. The isolated LECs can be cultured for up to four generations, after which they become senescent. They have been frozen, thawed, and successfully stimulated with angiopoietin-2. Due to the limited number of cell passages, we have no...

Watch this Scientific Journal Video about Murine Dermal Lymphatic Endothelial Cell Isolation at JoVE.com

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.

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 ...

Our group's research focuses on delineating the mechanisms regulating endothelial cell physiology. This protocol describes a method for magnetic bead-based isolation of renal endothelial cells, specifically from thermal lymphatic capillaries in facilities where fluorescence-activated cell sorting is not available. The protocol allows the isolation of lymphatic endothelial cells from transgenic mice to study their physiology in vitro.It is particularly suitable where the purity of the lymphatic endothelial cells can be compromised for downstream applications. This process provides a high yield of lymphatic endothelial cells obtained from the lymphatic capillaries and not other collecting vessels. It is also a nice alternative in the case of the absence of cell sorting facilities.

murine-dermal-lymphatic-endothelial-cell-isolation

Research

JoVE Journal

Biology

1.0K Views.  Texas Tech University Health Sciences Center. Our group's research focuses on delineating the mechanisms regulating endothelial cell physiology. This protocol describes a method for magnetic bead-based isolation of renal endothelial cells, specifically from thermal lymphatic capillaries in facilities where fluorescence-activated cell sorting is not available. The protocol allows the isolation of lymphatic endothelial cells from transgenic mice to study their physiology in vitro.It is particularly suitable where the purity of the l...

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

Isolation of Human Umbilical Vein Endothelial Cells (HUVEC)

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps include: degradation of the basement membrane, proliferation and migration (sprouting) of endothelial cells (EC) into the extracellular matrix, alignment of EC into cords, lumen formation, anastomosis, and formation of a new basement membrane. Many in vitro assays have been developed to study this process, but most only mimic certain stages of angiogenesis, and morphologically the vessels often do not resemble vessels in vivo. Here we demonstrate an optimized in vitro angiogenesis assay that utilizes human umbilical vein EC and fibroblasts. This model recapitulates all of the key early stages of angiogenesis, and importantly the vessels display patent intercellular lumens surrounded by polarized EC. Vessels can be easily observed by phase-contrast and time-lapse microscopy, and recovered in pure form for downstream applications.

Isolation of Human Umbilical Vein Endothelial Cells HUVEC

This video protocol illustrates the isolation and culture of human umbilical vein endothelial cells (HUVEC) from human umbilical cord. Once isolated these cells can be used for in vitro angiogenesis assays like the Optimized Fibrin Gel Bead Assay also demonstrated by the Hughes lab.

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps ...

Murine Pancreatic Islet Isolation

Isolation and Large Scale Expansion of Adult Human Endothelial Colony Forming Progenitor Cells

This paper introduces a novel recovery strategy for endothelial colony forming progenitor cells (ECFCs) from heparinized but otherwise unmanipulated adult human peripheral blood within a mean of 12 days. After large scale expansion &gt;1x108 ECFCs can be obtained for further tests. Advantageously by using pHPL the contact of human cells with bovine serum antigens can be excluded. By flow cytometry and immunohistochemistry the isolated cells can be characterized as ECFC and their in vitro functionality to form vascular like structures can be tested in a matrigel assay. Further these cells can be subcutaneously injected in a mouse model to form functional, perfused vessels in vivo. After long term expansion and cryopreservation proliferation, function and genomic stability appear to be preserved. 3,4
This animal-protein free isolation and expansion method is easily applicable to generate a large quantity of ECFCs.

Endothelial colony forming progenitor cells (ECFCs) are a promising tool to study vascular homeostasis and repair.1,2 This paper introduces a novel animal-serum free method for isolation and expansion of ECFC from heparinised adult human peripheral blood with pooled human platelet lysate (pHPL) diminishing the risk of anti-bovine immunisation.

This paper introduces a novel recovery strategy for endothelial colony forming progenitor cells (ECFCs) from heparinized but otherwise unmanipulated adult ...

Isolation and Animal Serum Free Expansion of Human Umbilical Cord Derived Mesenchymal Stromal Cells (MSCs) and Endothelial Colony Forming Progenitor Cells (ECFCs)

The umbilical cord is a rich source for progenitor cells with high proliferative potential including mesenchymal stromal cells (also termed mesenchymal stem cells, MSCs) and endothelial colony forming progenitor cells (ECFCs). Both cell types are key players in maintaining the integrity of tissue and are probably also involved in regenerative processes and tumor formation.
To study their biology and function in a comparative manner it is important to have both cells types available from the same donor. It may also be beneficial for regenerative purposes to derive MSCs and ECFCs from the same tissue.
Because cellular therapeutics should eventually find their way from bench to bedside we established a new method to isolate and further expand progenitor cells without the use of animal protein. Pooled human platelet lysate (pHPL) replaced fetal bovine serum in all steps of our protocol to completely avoid contact of the cells to xenogeneic proteins.
This video demonstrates a methodology for the isolation and expansion of progenitor cells from one umbilical cord.
All materials and procedures will be described.

Isolation and Animal Serum Free Expansion of Human Umbilical Cord Derived Mesenchymal Stromal Cells MSCs and Endothelial Colony Forming Progenitor Cells ECFCs

This protocol describes the isolation and subsequent expansion of mesenchymal stromal cells and endothelial colony forming cells without the use of animal serum to generate autologous pairs for experimental transplantation purposes.

The umbilical cord is a rich source for progenitor cells with high proliferative potential including mesenchymal stromal cells (also termed mesenchymal ...

Isolation of Valvular Endothelial Cells

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system. These thin, fibrous tissues are subjected to significant mechanical stresses as they open and close several billion times over a lifespan. The incredible endurance of these tissues is due to the resident valvular endothelial (VEC) and interstitial cells (VIC) that constantly repair and remodel in response to local mechanical and biological signals. Only recently have we begun to understand the unique behaviors of these cells, for which in vitro experimentation has played a key role. Particularly challenging is the isolation and culture of VEC. Special care must be used from the moment the tissue is removed from the host through final plating. Here we present protocols for direct isolation, side specific isolation, culture, and verification of pure populations of VEC. We use enzymatic digestion followed by a gentle swab scraping technique to dislodge only surface cells. These cells are then collected into a tube and centrifuged into a pellet. The pellet is then resuspended and plated into culture flasks pre-coated with collagen I matrix. VEC phenotype is confirmed by contact inhibited growth and the expression of endothelial specific markers such as PECAM1 (CD31), Von Willebrand Factor (vWF), and negative expression of alpha-smooth muscle actin (&alpha;-SMA). The functional characteristics of VEC are associated with high levels of acetylated LDL. Unlike vascular endothelial cells, VEC have the unique capacity to transform into mesenchyme, which normally occurs during embryonic valve formation1. This can also occur during significantly prolonged post confluent in vitro culture, so care should be made to passage at or near confluence. After VEC isolation, pure populations of VIC can then be easily acquired.

We provide a method for isolating and culturing pure populations of heart valve endothelial cells (VEC). VEC can be isolated from either side of the cusp or leaflet and immediately following, underlying interstitial cell (VIC) isolation is straightforward.

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system.  These thin, fibrous tissues are ...

Isolation of Embryonic Ventricular Endothelial Cells

Cell culture has greatly enhanced our ability to assess individual populations of cells under myriad culture conditions. While immortalized cell lines offer significant advantages for their ease of use, these cell lines are unavailable for all potential cell types. By isolating primary cells from a specific region of interest, particularly from a transgenic mouse, more nuanced studies can be performed. The basic technique involves dissecting the organ or partial organ of interest (e.g. the heart or a specific region of the heart) and dissociating the organ to single cells. These cells are then incubated with magnetic beads conjugated to an antibody that recognizes the cell type of interest. The cells of interest can then be isolated with the use of a magnet, with a short trypsin incubation dissociating the cells from the beads. These isolated cells can then be cultured and analyzed as desired. This technique was originally designed for adult mouse organs but can be easily scaled down for use with embryonic organs, as demonstrated herein. Because our interest is in the developing coronary vasculature, we wanted to study this population of cells during specific embryonic stages. Thus, the original protocol had to be modified to be compatible with the small size of the embryonic ventricles and the low potential yield of endothelial cells at these developmental stages. Utilizing this scaled-down approach, we have assessed coronary plexus remodeling in transgenic embryonic ventricular endothelial cells.

Primary cell culture is a useful technique for analyzing specific populations of cells, particularly from transgenic mouse embryos at specific developmental stages. Herein, embryonic ventricles are dissected and dissociated, and antibody-conjugated beads recognize and separate out the endothelial cells for further analysis.

Cell culture has greatly enhanced our ability to assess individual populations of cells under myriad culture conditions. While immortalized cell lines ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...

Isolation of Mouse Coronary Endothelial Cells

Endothelial cells line the inner wall of blood vessels and play an important role in the regulation of vascular tone, vascular permeability, and new vascular formation. Endothelial cell dysfunction is implicated in the development and progression of many cardiovascular diseases including ischemic heart disease. To examine the function and characterization of coronary endothelial cells, cell isolation is the first step and it requires high purity and quantity to conduct subsequent experiments. This protocol describes an efficient method to isolate adult mouse coronary endothelial cells. The mouse heart is dissected and minced into small pieces. After the digestion of the heart using dispase and collagenase II, cells are washed&#160;and incubated with magnetic beads which are conjugated with anti-CD31 antibody. The beads with endothelial cells are washed several times and are ready to use in various applications, including imaging and molecular biological experiments. Efficient isolation yields approximately 104 cells per one heart with over 90% purity.

This protocol is prepared to share our method of isolating mouse coronary endothelial cells for the purpose of imaging or to conduct molecular biological experiments.

Endothelial cells line the inner wall of blood vessels and play an important role in the regulation of vascular tone, vascular permeability, and new ...

Quantifying Leukocyte Egress via Lymphatic Vessels from Murine Skin and Tumors

Leukocyte egress from peripheral tissues to draining lymph nodes is not only critical for immune surveillance and initiation but also contributes to the resolution of peripheral tissue responses. While a variety of methods are used to quantify leukocyte egress from non-lymphoid, peripheral tissues, the cellular and molecular mechanisms that govern context-dependent egress remain poorly understood. Here, we describe the use of in situ photoconversion for quantitative analysis of leukocyte egress from murine skin and tumors. Photoconversion allows for the direct labeling of leukocytes resident within cutaneous tissue. Though skin exposure to violet light induces local inflammatory responses characterized by leukocyte infiltrates and vascular leakiness, in a head-to-head comparison with transdermal application of fluorescent tracers, photoconversion specifically labeled migratory dendritic cell populations and simultaneously enabled the quantification of myeloid and lymphoid egress from cutaneous microenvironments and tumors. The mechanisms of leukocyte egress remain a missing component in our understanding of intratumoral leukocyte complexity, and thus the application of the tools described herein will provide unique insight into the dynamics of tumor immune microenvironments both at steady state and in response to therapy.

Here, we demonstrate the methods for in vivo quantification of leukocyte egress from na&#239;ve, inflamed, and malignant murine skin. We perform a head-to-head comparison of two models: transdermal FITC application and in situ photoconversion. Furthermore, we demonstrate the utility of photoconversion for tracking leukocyte egress from cutaneous tumors.

Leukocyte egress from peripheral tissues to draining lymph nodes is not only critical for immune surveillance and initiation but also contributes to the ...

Microvascular and Macrovascular Endothelial Cell Isolation and Purification from Lung-Derived Samples

The availability of cells isolated from healthy and diseased tissues and organs represents a key element for personalized medicine approaches. Although biobanks can provide a wide collection of primary and immortalized cells for biomedical research, these do not cover all experimental needs, particularly those related to specific diseases or genotypes. Vascular endothelial cells (ECs) are key components of the immune inflammatory reaction and, thus, play a central role in the pathogenesis of a variety of disorders. Notably, ECs from different sites display different biochemical and functional properties, making the availability of specific EC types (i.e., macrovascular, microvascular, arterial, and venous) essential for designing reliable experiments. Here, simple procedures to obtain high-yield, virtually pure human macrovascular and microvascular endothelial cells from the pulmonary artery and lung parenchyma are illustrated in detail. This methodology can be easily reproduced at a relatively low cost by any laboratory to achieve independence from commercial sources and obtain EC phenotypes/genotypes that are not yet available.

Although challenging, the isolation of pulmonary endothelial cells is essential for studies on lung inflammation. The present protocol describes a procedure for the high-yield, high-purity isolation of macrovascular and microvascular endothelial cells.

The availability of cells isolated from healthy and diseased tissues and organs represents a key element for personalized medicine approaches. Although ...