The authors would like to acknowledge the Baker Foundation and the Royal Children's Foundation 'Women in Science Fellowship' support of Zerina Lokmic. Andrew G. Elefanty and Edouard G. Stanley are NHMRC Senior Research Fellows. Work in their laboratories was supported by NHMRC and Stem Cells Australia.

Ethics statement: Ethical approval for collection of lymphatic malformation and foreskin tissues was obtained from the Human Research Ethics Committees at the Royal Children&#8217;s Hospital, Melbourne, Australia. Signed consent was received from patients&#8217; parents prior to surgery. Tissue samples were collected from patients diagnosed with LMs undergoing surgical procedures as part of their clinical management and patients undergoing elective circumcision. All experiments were performed in accordance with guidelines of the National Health and Medical Research Council, Australia.
1. Preparation of Cell Suspension from Foreskin and Lymphatic Malformation Tissues
<ol>
	<li>Buffers and Media Preparation.
	<ol>
		<li>Prepare complete endothelial cell media using commercially available EGM-2 MV Bullet Kit by warming EGM-2 media and gently thawing the kit components in 37 &#176;C water bath. In class II biosafety cabinet, aseptically add each component to the EGM-2 media. Once all components are added to the media, refer to this media as &#8216;complete endothelial cell media&#8217;. Use sterile 50 ml pipette to mix the contents prior to use. 
		NOTE: All steps pertaining to cell culture are performed in a class II biohazard cabinet. All solutions are stored at 4 &#176;C as per manufacturers&#8217; instructions and are warmed to room temperature prior to use. The complete endothelial cell media, cell culture buffers and enzyme solutions are warmed at 37 &#176;C for 20 min prior to use.</li>
		<li>Supplement complete endothelial cell media with 50 ng/ml VEGF-C. Aliquot the supplemented endothelial cell media into 50 ml sterile tubes and store at 4 &#176;C until use. The media is stable for at least 4 weeks when stored at 4 &#176;C.</li>
		<li>Prepare calcium and magnesium free phosphate-buffered saline (PBS) by dissolving 8.752 g NaCl, 1.416 g Na2HPO4.2H2O and 0.395 g KH2PO4 in 1,000 ml of water. Adjust pH to 7.4. Filter sterilize PBS using 0.22 &#956;m filter and store at 4 &#176;C. Prior to use add antibiotic/antimycotic solution (used at 1:100).</li>
		<li>To prepare human fibronectin, dissolve 1 mg of fibronectin in 10 ml of sterile water. Store reconstituted solution in 100 &#956;l aliquots at -20 &#176;C. On the day of use add 10 ml of sterile PBS to 100 &#956;l of fibronectin aliquot (to give a final working concentration of 10 &#956;g/ml).</li>
		<li>For the enzyme media, prepare a solution containing 0.04% Dispase II, 0.25% Collagenase II and 0.01% DNase I in sterile PBS. First, weigh dispase and collagenase, place in a sterile 50 ml tube, add the required volume of PBS and incubate for 30 min with shaking at 37 &#176;C to dissolve. Once dissolved, filter sterilize (0.22 &#956;m filter) dispase/collagenase solution in biosafety hood. Aseptically add the DNase I. Store the enzymatic solution at 37 &#176;C until use. 
		NOTE: DNase I is a commercial cell culture grade reagent available as sterile lyophilized powder.</li>
	</ol>
	</li>
</ol>
2. Preparation of Cell Suspension from Foreskin and Lymphatic Malformations
<ol>
	<li>Weigh a 50 ml tube containing 10 ml of DMEM/2% antibiotic antimycotic solution. This tube will be used for tissue collection.</li>
	<li>Following surgical removal of lymphatic malformation and foreskin tissues, aseptically add tissue specimen to the tube and transfer on ice to the cell culture laboratory.</li>
	<li>Weigh the tube containing the tissue and calculate tissue weight. Use this weight to calculate the volume of enzymatic solution needed to digest the tissue.</li>
	<li>In a Class II biosafety cabinet, use sterile forceps to transfer the tissue and media into a 100 mm tissue culture dish. Use sterile scissors to finely mince the tissue into ~1 mm3 pieces.</li>
	<li>Transfer minced tissue mixed with media into a 50 ml tube containing an additional 10 ml of sterile DMEM/2% antibiotic antimycotic solution. Mix by inversion to re-suspend the tissue. Centrifuge the sample at 300 x g for 5 min.</li>
	<li>Remove the supernatant. Based on tissue weight, add 1 ml of pre-warmed (37 &#176;C) collagenase II/DNase I/ Dispase II digestion solution per 100 mg of minced tissue. Generally, use 10 ml of enzyme solution for approximately 1 g of minced tissue.</li>
	<li>Incubate the tissue in a 37 &#176;C incubator for 20-90 min with constant shaking at 200 rpm. At the end of this period, ensure that nearly all of the tissue is digested into fine fragments. 
	NOTE: Exposure of the cells to collagenase and dispase at the time of isolation from the primary tissues influences cell survival. We have found empirically that most neonatal foreskin samples optimally require 20-30 min of digestion, whereas fibrotic lymphatic malformation samples may require 60-90 min of digestion. LM LEC yields are influenced by the amount of fibrosis present, more fibrous tissue yielding fewer cells, possibly due to deleterious effects of prolonged exposure to collagenase II and dispase II.</li>
	<li>Following digestion, transfer the tube to the biosafety cabinet and pass the digested tissue solution through a 70 &#956;m strainer placed into a sterile 50 ml tube. Using 3 ml syringe piston with rubber plunger, grind down the remaining tissue until only small traces of extracellular matrix are observed.</li>
	<li>Wash the cell strainer with a volume of endothelial cell medium equivalent to the volume of dissociating enzyme medium to recover cells stuck to sieve and to inactivate the enzymes. For example, for 2 ml of dissociating enzyme medium to digest the minced tissue, add 2 ml of endothelial cell medium to inactivate the enzymes.</li>
	<li>Centrifuge the cell solution at 300 x g for 5 min and aspirate the supernatant.&#160;R>e-suspend the cells in 10 ml of PBS. Centrifuge cells at 300 x g for 5 min, remove the supernatant then repeat cell wash twice.&#160;>Resuspend the cell pellet in 20 ml of endothelial cell medium. Count cells using trypan blue then seed 2 x 10>6 cells in 150 cm>2 flask pre-coated with fibronectin in a final volume of 20 ml of endothelial cell medium. Culture at 37 &#176;C in a 5% CO>2 in humidified air incubator.> 
	NOTE: It is expected that 75%-90% of nucleated cells survive tissue digestion as estimated by trypan blue exclusion. The initial cell count reflects all nucleated cells in the cell suspension (i.e. keratinocytes, leukocytes, macrophages, connective tissue cells and blood vessel cells). The cell count at 5-7 days reflects cells that have attached, survived and proliferated during cell culture. Coating tissue culture flasks with fibronectin also improves subsequent LEC and LM LEC survival.</li>
	<li>After overnight incubation, wash away unbound cells in three washes with PBS/1% antibiotic-antimycotic solution and add fresh endothelial cell medium. Carry out three washes to effectively remove unbound cells and red blood cells present in the flask. Change media every second day. Cells will be ~80% confluent after 5-7 days.</li>
</ol>
3. Antibody Staining of Cells for Lymphatic Endothelial Cell Surface Markers for Flow Cytometry
<ol>
	<li>After the cells have reached 80% confluence, aspirate medium and rinse cells with PBS.</li>
	<li>Detach adherent cells by incubating cells with 7 ml of cell detachment solution such as Accutase per 150 cm2 flask for 5-7 min at 37 &#176;C.</li>
	<li>Harvest detached cells, inactivate the cell detachment solution by adding 3 volumes of endothelial cell medium and transfer the cell suspension to a new sterile tube.</li>
	<li>Centrifuge the cells at 300 x g for 5 min.</li>
	<li>Aspirate supernatant and re-suspend the cells in 5 ml of PBS. Centrifuge cells at 300 x g for 5 min, remove the supernatant then repeat cell wash. Re-suspend the cells in 5 ml of PBS.</li>
	<li>Count the number of viable cells. Mix 10 &#956;l of cell suspension with 90 &#956;l of 0.4% trypan blue. Transfer 10 &#956;l of this suspension to hemocytometer for counting. 
	NOTE: The final cell yield will depend on the sample size processed. The usual cell yield per 150 cm2 flask ranges from 7 x 105 to 1.2 x 106 of viable cells.</li>
	<li>Prepare the antibody solutions for cell staining. 
	NOTE: The following dilutions were titrated for staining 1 x 106 in a staining volume of 100 &#956;l.
	<ol>
		<li>Dilute PE conjugated mouse anti human VEGFR-3 (1:50); PE-Cy7 conjugated mouse anti human conjugated CD34 (1:200); APC-conjugated mouse anti human CD31 (1:100) and Alexa 488-conjugated rat anti human PODOPLANIN (1:200) in total volume of 100 &#956;l sterile 5% FBS/PBS solution per tissue sample. Following preparation of antibody solution, keep on ice until use. Similarly prepare diluted isotype control antibody mixture to facilitate setting of flow cytometry gates.</li>
	</ol>
	</li>
	<li>To reduce nonspecific binding of the antibodies, centrifuge cells at 300 x g for 2 min, remove supernatant then suspend the cells in 100 &#956;l sterile 5% FBS/PBS solution per sample to be stained. Incubate cells on ice for 20 min at 4 &#176;C.</li>
	<li>Centrifuge cells at 300 x g for 5 min, remove supernatant then re-suspend cells in the conjugated antibody cocktail. Also stain a smaller aliquot of cells (1 x 105) in 100 &#956;l isotype control antibody mixture. Incubate cells on ice for 20 min.</li>
	<li>To remove the unbound antibody, add 2 ml of 2% FBS/PBS solution and centrifuge the cells at 300 x g for 5 min. Aspirate the supernatant and repeat the wash.</li>
	<li>Resuspend the cell pellet for the isotype control and the antibody stained sample to be sorted in 300 &#956;l of 0.5 mg/ml propidium iodide/2% FBS/PBS solution. Place tubes on ice until sorting.</li>
</ol>
4. Cell Sorting
<ol>
	<li>Use the following materials to set up the instrument; alignment beads such as commercially available fluorescent particles, phosphate-buffered saline based sheath fluid and drop delay beads, such as Accudrop fluorescent beads.</li>
	<li>Isolate the purified human LECs on a multi-parameter fluorescence-activated cell sorting instrument. Set up the instrument and align as per manufacturer&#8217;s recommendations. In addition, run single stain compensation controls with each sort to generate the compensation matrix and to thus eliminate substantial bleed through of the emission of fluorochromes such as PE into the PE-Cy channel. 
	NOTE: A higher proportion of the FACS sorted cells survive the isolation if a 100 &#956;m nozzle is used and cells are sorted into endothelial cell medium.</li>
</ol>
5. Cell Culture Post FACS Sorting
<ol>
	<li>Following sorting, centrifuge cells at 300 x g for 5 min. Aspirate the supernatant and resuspend the cells in 5-10 ml of media (depending on the flask used to seed the cell). If less than 50,000 cells are sorted, the cells are cultured in fibronectin coated 25 cm2 flask.</li>
	<li>Otherwise culture the cells in 75 cm2 fibronectin-coated flask at 37 &#176;C in a 5% CO2, air humidified incubator. Change media after 2 days. 
	NOTE: Cell media is changed every second day because prolonging the time between media changes appears to favor survival of the non-endothelial cells. We validate the lymphatic endothelial cell phenotype using immunohistochemical detection of markers: PROX-1, VEGFR-3, Podoplanin, CD34 and CD31 when the cells are first split for further expansion15.</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contractile+physiology+of+lymphatics.">Contractile physiology of lymphatics.</a> Lymphat Res Biol. 7 (2), 87-96 (2009).</li><li>Witte, M. 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=Structure+function+relationships+in+the+lymphatic+system+and+implications+for+cancer+biology.">Structure function relationships in the lymphatic system and implications for cancer biology.</a> Cancer Metastasis Rev. 25 (2), 159-184 (2006).</li><li>Andrade, M., Jacomo, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomy+of+the+human+lymphatic+system.">Anatomy of the human lymphatic system.</a> Cancer Treat Res. 135, 55-77 (2007).</li><li>Kumar, V., Abbas, A. 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The authors have nothing to disclose.

LECs play an important role in maintaining fluid homeostasis, immune response to foreign antigens and absorption and transport of some nutrients. LEC homeostasis can be affected by disease processes such as bacterial infections and tumor metastasis, but LECs can also develop somatic mutations that result in formation of dysfunctional lymphatic vessels and significant morbidity for affected patients. To gain more understanding of lymphatic malformation etiology through in vivo implantation and ex vivo ex...

A major function of the lymphatic vascular system is to absorb lymph, an excess interstitial fluid containing lipids, proteins and cellular components, and conduct it to the blood venous system. A network of lymphatic capillaries directs lymph to the lymph nodes where it is screened for presence of foreign antigens, an important process in immune surveillance and deployment of white blood cells to neutralize foreign antigens.
The uni-directional lymphatic system starts in tissues with the initial lymphatic capillary, a unique structure with a discontinuous single layer of thin-walled flat endothelial cells with specialized cell junctions that permit lymph entry1,2. These capillaries are attached to the neighboring connective tissue matrix via anchoring filaments to prevent vessel collapse in presence of increased interstitial pressure3. The initial lymphatic capillaries empty into collecting lymphatic capillaries that coalesce into larger lymphatic vessels or veins. In comparison to initial lymphatic capillary vessels, collecting lymphatic vessels have thicker vessel walls, paired lymphatic valves and are encased by a discontinuous basement membrane in which a few smooth muscle cells are embedded4. Coordinated opening and closure of lymphatic valves and contraction of smooth muscle cells facilitates flow of lymph3. In humans, the lymphatic veins from various regions of the body join to form lymphatic trunks which merge to form two lymphatic ducts: the thoracic duct and the right lymphatic duct. The thoracic duct drains lymph from the left side of the body and from the right side below the chest while the right lymphatic duct drains lymph from the right arm and right side of the head, neck, and thorax. Both ducts conduct lymph into the subclavian veins in the neck5.
Disorders of the lymphatic system are broadly grouped into acquired and congenital (Table 1). Examples of acquired conditions are lymphangitis and secondary lymphedema. Lymphangitis is an inflammation of a lymphatic vessel due to bacterial infection. The affected lymphatics dilate and fill with exudate containing polymorphonuclear cells. In skin, these lymphatics are visible as red, painful subcutaneous streaks often accompanied by enlargement of the associated draining lymph node (lymphadenitis)6. Secondary lymphedema arises as a consequence of damage or obstruction to the lymphatic vessel or lymph node obstruction. This leads to chronic progressive swelling due to accumulation of lymph distal to the damage or obstruction. In developed countries, secondary lymphedema is most commonly associated with malignancy where metastasizing tumors obstruct lymphatic vessels or regional lymph nodes, or as a consequence of anti-cancer therapy following surgical removal of lymph nodes, post-irradiation fibrosis and post-inflammatory thrombosis and scarring7. In other parts of the world, secondary lymphedema may be secondary to lymphatic obstruction caused by parasitic worms such as Wuchereria bancrofti6.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td colspan="4">Disorders of Lymphatic Vascular System</td>
		</tr>
		<tr>
			<td>Acquired</td>
			<td colspan="3">Congenital</td>
		</tr>
		<tr>
			<td rowspan="3">Lymphadenitis 
			Secondary lymphedema</td>
			<td>Primary Lymphedema10</td>
			<td>Sporadic Lymphatic Malformations13</td>
			<td>Lymphatic Malformations Associated with Syndromes13</td>
		</tr>
		<tr>
			<td rowspan="2">e.g. 
			Milroy Syndrome 
			Meige Syndrome</td>
			<td>Simple: 
			Lymphatic malformations</td>
			<td rowspan="2">e.g. 
			Klippel-Tranaunay Syndrome 
			Parks Weber Syndrome 
			Sturge-Weber Syndrome</td>
		</tr>
		<tr>
			<td>Combined: 
			Capillary-lymphatic malformations 
			Capillary-lymphatic-venous malformation 
			Capillary-lymphatic-arteriovenous malformation 
			Capillary-lymphatic venous-arteriovenous malformation</td>
		</tr>
	</tbody>
</table>
Table 1. Overview of the disorders of lymphatic vascular system.
Congenital disorders of the lymphatic system include primary (idiopathic) lymphedema thought to be caused by genetic mutations, lymphangiectasia and anomalies of the lymphatic system8,9. Primary lymphedema can be sporadic presumably caused by de novo mutations, or inherited. Lymphatic disorders can also be isolated or comprise part of a more generalized syndrome10. In the pediatric population, 97% of lymphedema is sporadic with abnormalities in lymphatic vessel structure that impair regional lymph drainage11. Milroy disease is an example of primary lymphedema caused by mutation in the VEGFR-3 gene evident at birth or soon after12. Although mostly familial condition, the Milroy disease can also be identified in infants without family history of Milroy disease32. The severity of any lymphedema is dependent on the amount of lymph production and ability to transport lymph back to venous circulation6.
Based on clinical presentation and in situ endothelial cell proliferation, anomalies of the lymphatic system are classified as lymphatic tumors or lymphatic malformations13. Kaposiform lymphangiomatosis is an example of an LEC tumor14. Lymphatic malformations are thought to arise during embryonic development and grow in proportion to the child15,16. They rarely regress but can remain asymptomatic until trauma or infection precipitates rapid growth leading to clinical complications. The orderly structure of lymphatic network and conduction of lymph from the tissue to venous circulation described above is perturbed in lymphatic malformations which consist of localized collections of abnormal cystic structures filled with lymphatic fluid. While there is no clinical or experimental evidence that these cystic vessels are connected to the lymphatic circulation or that they contain functional lymphatic valves, their lymphatic identity is confirmed by expression of range of lymphatic cell markers such as PODOPLANIN, CD31, Lymphatic Vessel Endothelial Receptor 1 (LYVE-1), Prospero homeobox protein 1 (PROX-1) and VEGFR-315,17,18. These cystic structures can be either small (microcystic) or large (macrocystic), but most lymphatic malformations contain both microcystic and macrocystic components (Figure 1)16. Following surgery, injection sclerotherapy and/or radiofrequency ablation the lymphatic malformations often reoccur.
<img alt="Figure 1" src="/files/ftp_upload/52691/52691fig1highres.jpg" width="700" /> 
Figure 1. Morphology of human lymphatic vessels and lymphatic malformations. Normal human lymphatic (A) and lymphatic malformation vessels (B and C) labelled with antibody to PODOPLANIN (brown label, arrow). Human lymphatic malformation vessels are characterized by marked dilation and considerable variation in lumen size. These localized abnormal cystic structures can be either small (microcystic, *) (B) or large (macrocystic, #) (C). Most lymphatic malformations contain both microcystic and macrocystic components. <a href="https://www.jove.com/files/ftp_upload/52691/52691fig1highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>
Some investigators have suggested that lymphatic malformations represent a developmental disorder of lymphatic vasculature in which the LECs do not have abnormal growth potential but instead have failed to connect to the normal circulation19. However, we have found that the LM LECs proliferate faster and are more resistant to apoptosis than foreskin LECs15 suggesting that there is a primary defect in the LM LECs. When LM LECs are implanted in a mouse xenograft model, they form structures reminiscent of lymphatic malformations15. This supports a hypothesis that lymphatic malformations may be caused by one or more somatic mutations arising in LM LECs during fetal development. Indeed, recent reports have identified one such mutation in the p110&#945; catalytic subunit of Phosphoinositide-3-Kinase (PIK3CA) gene20.
Given the advances in DNA sequencing technology, relevant mutations could be more readily identified in isolated LM LECs, guiding future studies of these conditions. The isolation of viable LECs would facilitate comparisons between abnormal and normal LECs in assays such as migration, proliferation, tube forming ability and survival in response to reduced nutrient availability or pro-apoptotic agents15. Isolated LECs would further enable us to perform cell-specific gene expression and proteomic studies, to delineate new LEC subpopulations and discover novel pharmacological agents suitable for clinical management of lymphatic malformations.
We have previously published a LEC isolation method based on magnetic bead separation of LECs from neonatal foreskin and lymphatic malformations15. We reported a strategy of separating normal and diseased LECs from vascular endothelial cells based on the absence of CD34 expression, followed by subjecting CD34Neg cell fraction to positive selection for CD31. However, this method was hampered by the presence of residual non-endothelial cells. This was independent of removing epidermis prior to subsequent connective tissue digestion. These contaminants generally proliferated more rapidly and thus eventually overgrew the endothelial cell cultures despite subsequent attempts to repeat LEC isolation. Indeed, an initial contamination of non-endothelial cells as low as 2% to 5% was sufficient to overwhelm the LEC population15. This prompted us to explore fluorescently activated cell sorting method as an option to improve LEC cell yield and purity. In addition, we used multi-parameter sorting to enhance the specificity of the LEC populations, adding VEGFR-3 and PODOPLANIN to the selection markers to identify CD34LowCD31PosVEGFR-3PosPODOPLANINPos LECs.
The rationale for selecting these markers was based on the reports that while LECs and blood vascular endothelial cells have many cell surface markers in common such as CD31, LECs show phenotypic variation in their expression of CD34, PODOPLANIN and VEGFR-3 cell surface marker when compared to blood vascular endothelial cells21-23. CD31 is a 130 kDa transmembrane glycoprotein also known as platelet endothelial cell adhesion molecule 1 (PECAM-1). It is considered to be a pan-endothelial cell marker since it is expressed on all types of blood and lymphatic vessels21,24,25. CD34 is 110-kDa transmembrane glycoprotein present on most hematopoietic progenitor and stem cells, vascular endothelial cells and some lymphatic vessels26.
VEGFR-3, the receptor for vascular endothelial growth factors C and D, is initially present on the developing veins in the mouse embryo, but following lymphatic specification regulated by the transcription factors SRY-related HMG-box (SOX)-18, chicken ovalbumin upstream promoter transcription factor 2 (COUPTF-II) and PROX-1, VEGFR-3 venous expression is lost and it becomes restricted to embryonic LECs25,27. PODOPLANIN, a 38 kDa membrane mucoprotein, is first noted on lymphatic vessels at approximately embryonic day 11 (~E11.0) of mouse embryonic development 28 and whilst it is strongly expressed by microvascular lymphatic vessels, PODOPLANIN expression by macrocystic lymphatic endothelium in lymphatic malformations is more variable15. Flow cytometry experiments suggest that at least some CD34HighCD31Pos endothelial cells express the lymphatic marker PODOPLANIN29. Although systematic evaluation of LYVE-1 and PODOPLANIN staining in human lymphatic malformations showed that both are effective at staining lymphatic malformation endothelium30, in normal tissues, LYVE-1 was reported to be strongly present in the initial lymphatic capillary endothelium but reduced and even absent in the collecting lymphatic endothelium31. As our aim is to isolate both the initial and collecting lymphatic endothelial cells we have opted not to use LYVE-1 as part of our cell selection strategy. Finally, the decision to employ these markers was also based on the availability of antibodies that are used diagnostically for labelling lymphatic vessels for microscopic imaging, a feature that would permit correlation between flow cytometry and immunofluorescent studies.
This article will describe the tissue digestion method, cell staining and FACS settings required for successful isolation of CD34LowCD31PosVEGFR-3PosPODOPLANINPos LECs as well as CD34HighCD31PosVEGFR-3PosPODOPLANINPos endothelial cells from foreskin and lymphatic malformation tissue.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Name of Material/Equipment</td>
			<td>Company</td>
			<td>Catalogue Number</td>
			<td>Description</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Life Technologies (Gibco)</td>
			<td>11965-092</td>
			<td>Used to collect tissue samples from the operating theatre.</td>
		</tr>
		<tr>
			<td>EGM-2 MV Bullet Kit</td>
			<td>Lonza</td>
			<td>CC-3202</td>
			<td>EGM-2 MV Bullet Kit contains 500 ml of EBM-2 media and human EGF, hydrocortisone, gentamycin (GA-1000), fetal bovine serum (FBS), VEGF, human FGF-b, R3-IGF-1 and ascorbic acid.</td>
		</tr>
		<tr>
			<td>VEGF-C</td>
			<td>R&#38;D Systems</td>
			<td>2179-VC-025</td>
			<td>Complete endothelial cell media is supplemented with 50 ng/mL VEGF-C</td>
		</tr>
		<tr>
			<td>Antibiotic-antimycotic solution (100x)</td>
			<td>Life Technologies (Gibco)</td>
			<td>15240-062</td>
			<td>This solution contains 10,000U/ml of penicillin, 10,000 mg/ml of streptomycin and 25 mg/ml of Fungizone antimycotic. Use at 1:100 dilution.</td>
		</tr>
		<tr>
			<td>Fibronectin from human plasma</td>
			<td>Sigma-Aldrich</td>
			<td>F2006</td>
			<td>Used at 10 mg/ml concentrations. It can be re-used for up to 1 month if kept at 4 degrees and maintained sterile. Use 7 mL to coat 150 cm2 flask.</td>
		</tr>
		<tr>
			<td>StemPro Accutase Cell dissociation reagent</td>
			<td>Life Technologies (Gibco)</td>
			<td>A11105-01</td>
			<td>StemPro&#174;Accutase&#174; is used at 0.05 ml per cm2 to cover the entire surface area of the flask. Using lesser volumes may result in incomplete cell detachment.</td>
		</tr>
		<tr>
			<td>0.4% Trypan Blue dye</td>
			<td>Life Technologies (Gibco)</td>
			<td>15250-061</td>
			<td>Used as a viability stain.</td>
		</tr>
		<tr>
			<td>Dispase II</td>
			<td>Roche Applied Biosciences</td>
			<td>4942078001</td>
			<td>Used at 0.04%</td>
		</tr>
		<tr>
			<td>Collagenase II</td>
			<td>Worthington Lab</td>
			<td>4176</td>
			<td>Used at 0.25%</td>
		</tr>
		<tr>
			<td>DNase I</td>
			<td>Roche Applied Biosciences</td>
			<td>11284932001</td>
			<td>Used at 0.01%</td>
		</tr>
		<tr>
			<td>70 mm nylon cell strainers</td>
			<td>BD Falcon</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>PE-conjugated VEGFR-3 clone 9D9F9, clone WM59</td>
			<td>BioLegend</td>
			<td>356204</td>
			<td>Used at 1:50 dilution</td>
		</tr>
		<tr>
			<td>PE-Cy7-conjugated CD34, clone 581</td>
			<td>BioLegend</td>
			<td>343516</td>
			<td>Used at 1:200 dilution</td>
		</tr>
		<tr>
			<td>APC-conjugated mouse anti human CD31, clone WM59</td>
			<td>BioLegend</td>
			<td>303116,</td>
			<td>Used at 1:100 dilution</td>
		</tr>
		<tr>
			<td>Alexa 488-conjugated rat anti human PODOPLANIN, clone NC-80.</td>
			<td>BioLegend</td>
			<td>337006</td>
			<td>Used at 1:200 dilution</td>
		</tr>
		<tr>
			<td>PE-conjugated mouse IgG1, k isotype, clone MOPC-21</td>
			<td>BioLegend</td>
			<td>400112</td>
			<td>Used at 1:50 dilution</td>
		</tr>
		<tr>
			<td>PE-Cy7-conjugated mouse IgG1, k isotype, clone MOPC-21</td>
			<td>BioLegend</td>
			<td>400126</td>
			<td>Used at 1:200 dilution</td>
		</tr>
		<tr>
			<td>APC-conjugated mouse IgG1, k isotype, clone MOPC-21</td>
			<td>BioLegend</td>
			<td>400120</td>
			<td>Used at 1:100 dilution</td>
		</tr>
		<tr>
			<td>Alexa 488-conjugated mouse IgG2a, k isotype, clone RATK2758</td>
			<td>BioLegend</td>
			<td>400525</td>
			<td>Used at 1:200 dilution</td>
		</tr>
	</tbody>
</table>

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.

Following initial tissue digestion, after 24 hours in culture of unfractionated samples, distinct endothelial cell colonies can be observed (Figure 2A) together with fibroblast-like cells and smooth muscle cells. Following sorting and after 24 hours in cell culture, the CD34LowCD31PosVEGFR-3PosPodoplaninPos cells attach and show typical cobblestone morphology (Figure 2B and C). Using the FACS method described above, we are able...

Watch this Scientific Journal Video about Isolation of Human Lymphatic Endothelial Cells by Multi-parameter Fluorescence-activated Cell Sorting at JoVE.com

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

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 ...

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14.2K Views.  The Royal Children&#146;s Hospital. 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.

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

A Real-time Electrical Impedance Based Technique to Measure Invasion of Endothelial Cell Monolayer by Cancer Cells

Metastatic dissemination of malignant cells requires degradation of basement membrane, attachment of tumor cells to vascular endothelium, retraction of endothelial junctions and finally invasion and migration of tumor cells through the endothelial layer to enter the bloodstream as a means of transport to distant sites in the host1-3. Once in the circulatory system, cancer cells adhere to capillary walls and extravasate to the surrounding tissue to form metastatic tumors4,5. The various components of tumor cell-endothelial cell interaction can be replicated in vitro by challenging a monolayer of human umbilical vein endothelial cells (HUVEC) with cancer cells. Studies performed with electron and phase-contrast microscopy suggest that the in vitro sequence of events fairly represent the in vivo metastatic process6. Here, we describe an electrical-impedance based technique that monitors and quantifies in real-time the invasion of endothelial cells by malignant tumor cells.
Giaever and Keese first described a technique for measuring fluctuations in impedance when a population of cells grow on the surface of electrodes7,8. The xCELLigence instrument, manufactured by Roche, utilizes a similar technique to measure changes in electrical impedance as cells attach and spread in a culture dish covered with a gold microelectrode array that covers approximately 80% of the area on the bottom of a well. As cells attach and spread on the electrode surface, it leads to an increase in electrical impedance9-12. The impedance is displayed as a dimensionless parameter termed cell-index, which is directly proportional to the total area of tissue-culture well that is covered by cells. Hence, the cell-index can be used to monitor cell adhesion, spreading, morphology and cell density.
The invasion assay described in this article is based on changes in electrical impedance at the electrode/cell interphase, as a population of malignant cells invade through a HUVEC monolayer (Figure 1). The disruption of endothelial junctions, retraction of endothelial monolayer and replacement by tumor cells lead to large changes in impedance. These changes directly correlate with the invasive capacity of tumor cells, i.e., invasion by highly aggressive cells lead to large changes in cell impedance and vice versa. This technique provides a two-fold advantage over existing methods of measuring invasion, such as boyden chamber and matrigel assays: 1) the endothelial cell-tumor cell interaction more closely mimics the in vivo process, and 2) the data is obtained in real-time and is more easily quantifiable, as opposed to end-point analysis for other methods.

This article describes an in vitro technique for monitoring cancer cells invading through a monolayer of endothelial cells. The data is acquired in real-time as a function of changes in impedance on the surface of electrodes at the well bottom.

Metastatic dissemination of malignant cells requires degradation of basement membrane, attachment of tumor cells to vascular endothelium, retraction of ...

Isolation, Characterization and Comparative Differentiation of Human Dental Pulp Stem Cells Derived from Permanent Teeth by Using Two Different Methods

Developing wisdom teeth are easy-accessible source of stem cells during the adulthood which could be obtained by routine orthodontic treatments. Human pulp-derived stem cells (hDPSCs) possess high proliferation potential with multi-lineage differentiation capacity compare to the ordinary source of adult stem cells1-8; therefore, hDPSCs could be the good candidates for autologous transplantation in tissue engineering and regenerative medicine. Along with these benefits, possessing the mesenchymal stem cells (MSC) features, such as immunolodulatory effect, make hDPSCs more valuable, even in the case of allograft transplantation6,9,10. Therefore, the primary step for using this source of stem cells is to select the best protocol for isolating hDPSCs from pulp tissue. In order to achieve this goal, it is crucial to investigate the effect of various isolation conditions on different cellular behaviors, such as their common surface markers &amp; also their differentiation capacity.
Thus, here we separate human pulp tissue from impacted third molar teeth, and then used both existing protocols based on literature, for isolating hDPSCs,11-13 i.e. enzymatic dissociation of pulp tissue (DPSC-ED) or outgrowth from tissue explants (DPSC-OG). In this regards, we tried to facilitate the isolation methods by using dental diamond disk. Then, these cells characterized in terms of stromal-associated Markers (CD73, CD90, CD105 &amp; CD44), hematopoietic/endothelial Markers (CD34, CD45 &amp; CD11b), perivascular marker, like CD146 and also STRO-1. Afterwards, these two protocols were compared based on the differentiation potency into odontoblasts by both quantitative polymerase chain reaction (QPCR) &amp; Alizarin Red Staining. QPCR were used for the assessment of the expression of the mineralization-related genes (alkaline phosphatase; ALP, matrix extracellular phosphoglycoprotein; MEPE &amp; dentin sialophosphoprotein; DSPP).14

The method described isolation and characterization of human Dental Pulp Stem Cells (hDPSCs) by using either enzymatic dissociation of pulp (DPSC-ED) or direct outgrowth of stem cells from pulp tissue explants (DPSC-OG). Then followed by in vitro comparative differentiation of both types of hDPSCs into odontoblasts.

Developing wisdom teeth are easy-accessible source of stem cells during the adulthood which could be obtained by routine orthodontic treatments. Human ...

Cell Death Associated with Abnormal Mitosis Observed by Confocal Imaging in Live Cancer Cells

Phenanthrene derivatives acting as potent PARP1 inhibitors prevented the bi-focal clustering of supernumerary centrosomes in multi-centrosomal human cancer cells in mitosis. The phenanthridine PJ-34 was the most potent molecule. Declustering of extra-centrosomes causes mitotic failure and cell death in multi-centrosomal cells. Most solid human cancers have high occurrence of extra-centrosomes. The activity of PJ-34 was documented in real-time by confocal imaging of live human breast cancer MDA-MB-231 cells transfected with vectors encoding for fluorescent &gamma;-tubulin, which is highly abundant in the centrosomes and for fluorescent histone H2b present in the chromosomes. Aberrant chromosomes arrangements and de-clustered &gamma;-tubulin foci representing declustered centrosomes were detected in the transfected MDA-MB-231 cells after treatment with PJ-34. Un-clustered extra-centrosomes in the two spindle poles preceded their cell death. These results linked for the first time the recently detected exclusive cytotoxic activity of PJ-34 in human cancer cells with extra-centrosomes de-clustering in mitosis, and mitotic failure leading to cell death. According to previous findings observed by confocal imaging of fixed cells, PJ-34 exclusively eradicated cancer cells with multi-centrosomes without impairing normal cells undergoing mitosis with two centrosomes and bi-focal spindles. This cytotoxic activity of PJ-34 was not shared by other potent PARP1 inhibitors, and was observed in PARP1 deficient MEF harboring extracentrosomes, suggesting its independency of PARP1 inhibition. Live confocal imaging offered a useful tool for identifying new molecules eradicating cells during mitosis.

The cytotoxic activity of the phenanthridine PJ-34 in cancer cells undergoing mitosis was documented in real time by live confocal imaging. PJ-34 eradicated human breast cancer MDA-MB-231 cells harboring extra-centrosomes in mitosis. Unlike normal bi-focal mitosis, the extra-centrosomes were not clustered in the two spindle poles in the presence of PJ-34.

Phenanthrene derivatives acting as potent PARP1 inhibitors prevented the bi-focal clustering of supernumerary centrosomes in multi-centrosomal human ...

Isolation, Culture, and Imaging of Human Fetal Pancreatic Cell Clusters

For almost 30 years, scientists have demonstrated that human fetal ICCs transplanted under the kidney capsule of nude mice matured into functioning endocrine cells, as evidenced by a significant increase in circulating human C-peptide following glucose stimulation1-9. However in vitro, genesis of insulin producing cells from human fetal ICCs is low10; results reminiscent of recent experiments performed with human embryonic stem cells (hESC), a renewable source of cells that hold great promise as a potential therapeutic treatment for type 1 diabetes. Like ICCs, transplantation of partially differentiated hESC generate glucose responsive, insulin producing cells, but in vitro genesis of insulin producing cells from hESC is much less robust11-17. A complete understanding of the factors that influence the growth and differentiation of endocrine precursor cells will likely require data generated from both ICCs and hESC. While a number of protocols exist to generate insulin producing cells from hESC in vitro11-22, far fewer exist for ICCs10,23,24. Part of that discrepancy likely comes from the difficulty of working with human fetal pancreas. Towards that end, we have continued to build upon existing methods to isolate fetal islets from human pancreases with gestational ages ranging from 12 to 23 weeks, grow the cells as a monolayer or in suspension, and image for cell proliferation, pancreatic markers and human hormones including glucagon and C-peptide. ICCs generated by the protocol described below result in C-peptide release after transplantation under the kidney capsule of nude mice that are similar to C-peptide levels obtained by transplantation of fresh tissue6. Although the examples presented here focus upon the pancreatic endoderm proliferation and &#946;&#160;cell genesis, the protocol can be employed to study other aspects of pancreatic development, including exocrine, ductal, and other hormone producing cells.

A protocol to isolate, culture, and image islet cell clusters (ICCs) derived from human fetal pancreatic cells is described.&#160; The method details the steps necessary to generate ICCs from tissue, culture as monolayers or in suspension as aggregates, and image for markers of proliferation and pancreatic cell fate decisions.

For almost 30 years, scientists have demonstrated that human fetal ICCs transplanted under the kidney capsule of nude mice matured into functioning ...

Studying Pancreatic Cancer Stem Cell Characteristics for Developing New Treatment Strategies

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor initiation, metastasis and resistance to radio- and chemotherapy. Here we describe a specific methodology for culturing primary human pancreatic CSCs as tumor spheres in anchorage-independent conditions. Cells are grown in serum-free, non-adherent conditions in order to enrich for CSCs while their more differentiated progenies do not survive and proliferate during the initial phase following seeding of single cells. This assay can be used to estimate the percentage of CSCs present in a population of tumor cells. Both size (which can range from 35 to 250 micrometers) and number of tumor spheres formed represents CSC activity harbored in either bulk populations of cultured cancer cells or freshly harvested and digested tumors 1,2. Using this assay, we recently found that metformin selectively ablates pancreatic CSCs; a finding that was subsequently further corroborated by demonstrating diminished expression of pluripotency-associated genes/surface markers and reduced in vivo tumorigenicity of metformin-treated cells. As the final step for preclinical development we treated mice bearing established tumors with metformin and found significantly prolonged survival. Clinical studies testing the use of metformin in patients with PDAC are currently underway (e.g., NCT01210911, NCT01167738, and NCT01488552). Mechanistically, we found that metformin induces a fatal energy crisis in CSCs by enhancing reactive oxygen species (ROS) production and reducing mitochondrial transmembrane potential. In contrast, non-CSCs were not eliminated by metformin treatment, but rather underwent reversible cell cycle arrest. Therefore, our study serves as a successful example for the potential of in vitro sphere formation as a screening tool to identify compounds that potentially target CSCs, but this technique will require further in vitro and in vivo validation to eliminate false discoveries.

Pancreatic cancer stem cells (CSCs) can be expanded in vitro using the anchorage-independent sphere culture technique, which represents a powerful tool to study CSC biology and can serve as the first step to develop novel CSC-targeting therapies. Here the methodology for expanding, analyzing and targeting of pancreatic CSCs is provided.

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor ...

Isolation and Culture Expansion of Tumor-specific Endothelial Cells

Freshly isolated tumor-specific endothelial cells (TEC) can be used to explore molecular mechanisms of tumor angiogenesis and serve as an in vitro model for developing new angiogenesis inhibitors for cancer. However, long-term in vitro expansion of murine endothelial cells (EC) is challenging due to phenotypic drift in culture (endothelial-to-mesenchymal transition) and contamination with non-EC. This is especially true for TEC which are readily outcompeted by co-purified fibroblasts or tumor cells in culture. Here, a high fidelity isolation method that takes advantage of immunomagnetic enrichment coupled with colony selection and in vitro expansion is described. This approach generates pure EC fractions that are entirely free of contaminating stromal or tumor cells. It is also shown that lineage-traced Cdh5cre:ZsGreenl/s/l reporter mice, used with the protocol described herein, are a valuable tool to verify cell purity as the isolated EC colonies from these mice show durable and brilliant ZsGreen fluorescence in culture.

We report a reliable method to isolate and culture primary tumor-specific endothelial cells from genetically engineered mouse models.

Freshly isolated tumor-specific endothelial cells (TEC) can be used to explore molecular mechanisms of tumor angiogenesis and serve as an in vitro model ...

Isolation and Culture of Primary Endothelial Cells from Canine Arteries and Veins

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study vasculogenesis, (tumor) angiogenesis, and atherosclerosis. The current standard for in vitro research on human endothelial cells (ECs) is the use of Human Umbilical Vein Endothelial Cells (HUVECs) and Human Umbilical Artery Endothelial Cells (HUAECs). For canine endothelial research, only one cell line (CnAOEC) is available, which is derived from canine aortic endothelium. Although currently not completely understood, there is a difference between ECs originating from either arteries or veins. For a more direct approach to in vitro functionality studies on ECs, we describe a new method for isolating Canine Primary Endothelial Cells (CaPECs) from a variety of vessels. This technique reduces the chance of contamination with fast-growing cells such as fibroblasts and smooth muscle cells, a problem that is common in standard isolation methods such as flushing the vessel with enzymatic solutions or mincing the vessel prior to digestion of the tissue containing all cells. The technique we describe was optimized for the canine model, but can easily be utilized in other species such as human.

Novel isolation methods of primary endothelial cells from blood vessels are needed. This protocol describes a new technique that completely inverts blood vessels of interest, exposing only the endothelial side to enzymatic digestion. The resulting pure endothelial cell culture can be used to study cardiovascular diseases, disease modelling, and angiogenesis.

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study ...

Isolation of Endothelial Progenitor Cells from Healthy Volunteers and Their Migratory Potential Influenced by Serum Samples After Cardiac Surgery

Endothelial progenitor cells (EPCs) are recruited from the bone marrow under pathological conditions like hypoxia and are crucially involved in the neovascularization of ischemic tissues. The origin, classification and characterization of EPCs are complex; notwithstanding, two prominent sub-types of EPCs have been established: so-called &#34;early&#34; EPCs (subsequently referred to as early-EPCs) and late-outgrowth EPCs (late-EPCs). They can be classified by biological properties as well as by their appearance during in vitro culture. While &#34;early&#34; EPCs appear in less than a week after culture of peripheral blood-derived mononuclear cells in EC-specific media, late-outgrowth EPCs can be found after 2-3 weeks. Late-outgrowth EPCs have been recognized to be directly involved in neovascularization, mainly through their ability to differentiate into mature endothelial cells, whereas &#34;early&#34; EPCs express various angiogenic factors as endogenous cargo to promote angiogenesis in a paracrine manner. During myocardial ischemia/reperfusion (I/R), various factors control the homing of EPCs to regions of blood vessel formation.
Macrophage migration inhibitory factor (MIF) is a chemokine-like pro-inflammatory and ubiquitously expressed cytokine and was recently described to function as key regulator of EPCs migration at physiological concentrations1. Interestingly, MIF is stored in intracellular pools and can rapidly be released into the blood stream after several stimuli (e.g. myocardial infarction). 
This protocol describes a method for the reliable isolation and culture of early-EPCs from adult human peripheral blood based on CD34-positive selection with subsequent culture in medium containing endothelial growth factors on fibronectin-coated plates for use in in vitro migration assays against serum samples of cardiac surgical patients. Furthermore, the migratory influence of MIF on chemotaxis of EPCs compared to other well-known angiogenesis-stimulating cytokines is demonstrated.

Endothelial progenitor cells (EPCs) are crucially involved in the neovascularization of ischemic tissues. This method describes the isolation of human EPCs from peripheral blood, as well as the identification of their migratory potential against serum samples of cardiac surgical patients.

Endothelial progenitor cells (EPCs) are recruited from the bone marrow under pathological conditions like hypoxia and are crucially involved in the ...

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene and cell therapies are highly promising alternative approaches for CVD treatment. However, the broad clinical application of gene therapy is greatly limited by the lack of suitable gene delivery systems. The development of appropriate gene delivery vectors can provide a solution to current challenges in cell therapy. In particular, existing drawbacks, such as limited efficiency and low cell retention in the injured organ, could be overcome by appropriate cell engineering (i.e., genetic) prior to transplantation. The presented protocol describes the efficient and safe transient modification of endothelial cells using a polyethyleneimine superparamagnetic magnetic nanoparticle (PEI/MNP)-based delivery vector. Also, the algorithm and methods for cell characterization are defined. The successful intracellular delivery of microRNA (miR) into human umbilical vein endothelial cells (HUVECs) has been achieved without affecting cell viability, functionality, or intercellular communication. Moreover, this approach was proven to cause a strong functional effect in introduced exogenous miR. Importantly, the application of this MNP-based vector ensures cell magnetization, with accompanying possibilities of magnetic targeting and non-invasive MRI tracing. This may provide a basis for magnetically guided, genetically engineered cell therapeutics that can be monitored non-invasively with MRI.

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

This manuscript describes the efficient, non-viral delivery of miR to endothelial cells by a PEI/MNP vector and their magnetization. Thus, in addition to genetic modification, this approach allows for magnetic cell guidance and MRI detectability. The technique can be used to improve the characteristics of therapeutic cell products.

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene ...

Isolation and Profiling of Human Primary Mesenteric Arterial Endothelial Cells at the Transcriptome Level

Endothelial cells (ECs) are crucial for vascular and whole-body function through their dynamic response to environmental cues. Elucidating the transcriptome and epigenome of ECs is paramount to understanding their roles in development, health, and disease, but is limited in the availability of isolated primary cells. Recent technologies have enabled the high-throughput profiling of EC transcriptome and epigenome, leading to the identification of previously unknown EC cell subpopulations and developmental trajectories. While EC cultures are a useful tool in the exploration of EC function and dysfunction, the culture conditions and multiple passages can introduce external variables that alter the properties of native EC, including morphology, epigenetic state, and gene expression program. To overcome this limitation, the present paper demonstrates a method of isolating human primary ECs from donor mesenteric arteries aiming to capture their native state. ECs in the intimal layer are dissociated mechanically and biochemically with the use of particular enzymes. The resultant cells can be directly used for bulk RNA or single-cell RNA-sequencing or plated for culture. In addition, a workflow is described for the preparation of human arterial tissue for spatial transcriptomics, specifically for a commercially available platform, although this method is also suitable for other spatial transcriptome profiling techniques. This methodology can be applied to different vessels collected from a variety of donors in health or disease states to gain insights into EC transcriptional and epigenetic regulation, a pivotal aspect of endothelial cell biology.

The protocol describes the isolation, culture, and profiling of endothelial cells from human mesenteric artery. Additionally, a method is provided to prepare human artery for spatial transcriptomics. Proteomics, transcriptomics, and functional assays can be performed on isolated cells. This protocol can be repurposed for any medium- or large-size artery.