The authors wish to acknowledge funding from NIH/NHLBI R00 HL129068.

All animal studies were approved by the respective Institutional Animal Care and Use Committees (IACUC) at the Medical College of Wisconsin and Mayo Clinic.
NOTE: In this study, Yorkshire/Landrace/Duroc cross domestic pigs (Sus domesticus), male and female, 40-80 kg, 3-6 months old, were used.
1. Collection of porcine peripheral blood
<ol>
	<li>Prepare materials.
	<ol>
		<li>Dilute heparin solution to 100 U/mL in sterile saline.</li>
		<li>Add 3-4 mL of heparin solution to each of two 50 mL conical tubes and two 60 mL syringes.</li>
		<li>Draw undiluted heparin solution (1,000 U/mL) into an extension tube.</li>
		<li>Connect a 19 G needle to one end of the heparin-filled extension tube and a heparin-containing 60 mL syringe to the other end.</li>
	</ol>
	</li>
	<li>Anesthetize a pig according to Institutional policies
	<ol>
		<li>Administer an IM injection of atropine (0.05 mg/kg), tiletamine/zolazepam (2-5 mg/kg), and xylazine (2 mg/kg) to induce anesthesia. 
		NOTE: adequate anesthesia is achieved once the animal is unconscious and the mandibular muscles are non-rigid.</li>
		<li>Lay the animal in the supine position and place rolled-up towels on both sides to provide stability.</li>
		<li>Apply an ophthalmic ointment to the eyes to prevent dryness while under anesthesia.</li>
		<li>Provide thermal support during anesthesia including blankets, heating pads, and/or heated procedure table.</li>
	</ol>
	</li>
	<li>Clean the femoral groin area with betadine scrub solution.</li>
	<li>Puncture the femoral vein or artery with the 19 G needle and slowly (~1-2 mL/s) draw 50 mL of blood into the syringe. 
	NOTE: Drawing too fast can damage the cells.</li>
	<li>Leaving the needle in place, immediately kink the extension tube and disconnect the 60 mL syringe</li>
	<li>Slowly transfer the blood to a heparin-containing 50 mL conical tube.</li>
	<li>Tightly cap the 50 mL conical tube, gently invert a few times to mix, and place on ice.</li>
	<li>Connect the remaining heparin-containing 60 mL syringe to the extension tube, unkink the extension tube, and slowly draw an additional 50 mL of blood into the syringe.</li>
	<li>Immediately remove the needle from the artery or vein and slowly draw the remaining blood from the extension tube into the syringe.</li>
	<li>Disconnect the 60 mL syringe from the extension tube and slowly transfer the blood to the remaining heparin-containing 50 mL conical tube.</li>
	<li>Tightly cap the 50 mL conical tube, gently invert a few times to mix, and place on ice.</li>
	<li>Apply pressure hemostasis to the pig&#39;s femoral groin area.</li>
	<li>Recover the pig according to Institutional policies. Continuously monitor the animal until it regains sufficient consciousness to maintain sternal recumbency and return to the regular housing/kennel area.</li>
</ol>
2. Isolation of mononuclear cells
<ol>
	<li>Dilute the blood 1:1 with phosphate-buffered saline (PBS) in four 50 mL conical tubes. 
	NOTE: This work needs to be performed in a cell culture hood using an aseptic technique to avoid contaminating the cell culture.</li>
	<li>Add 25 mL of room temperature ready-to-use density gradient solution to eight 50 mL conical tubes.</li>
	<li>Very slowly pipette 25 mL of blood/saline solution along the inside of each density gradient solution containing 50 mL conical tube by holding the tube at a sharp angle to the pipette tip. 
	NOTE: The blood solution should gently layer on top of the density gradient solution and maintain a well-defined interface.</li>
	<li>Centrifuge the tubes for 30 min at 560 x g and room temperature (RT). 
	NOTE: For best results, disable the centrifuge brake if possible.</li>
	<li>Use a thin pipette to carefully collect the buffy coat (cloudy layer of mononuclear cells above the density gradient solution and below the clear plasma) from each tube and distribute evenly into two new 50 mL conical tubes. Discard the density gradient solution containing tubes. 
	&#8203;NOTE: Collect as much of the buffy coat as possible while collecting as little of the density gradient solution and plasma as possible.</li>
</ol>
3. Washing and plating of cells
<ol>
	<li>Coat a 6-well plate with fibronectin.
	<ol>
		<li>Make a stock solution of human plasma fibronectin by diluting it to 1 mg/mL in sterile water. Store the aliquots at -20 &#176;C.</li>
		<li>Make 3.6 mL of a working solution of fibronectin by diluting 600 &#181;L of fibronectin stock solution in 3 mL of PBS.</li>
		<li>Transfer 600 &#181;L of the fibronectin working solution into each well of a 6-well plate and gently rock until evenly coated.</li>
		<li>Wait 30 min for the fibronectin to coat the wells and gently aspirate the solution out of each well.</li>
	</ol>
	</li>
	<li>While waiting for the fibronectin to coat, wash the mononuclear cells
	<ol>
		<li>Top up the tubes with up to 45 mL of PBS</li>
		<li>Centrifuge for 5 min at 560 x g and 4 &#176;C with low brake. Aspirate the supernatant from both tubes.</li>
		<li>Resuspend each cell pellet in 25 mL of PBS.</li>
		<li>Repeat to wash a second time.</li>
	</ol>
	</li>
	<li>Resuspend each cell pellet in 5 mL of PBS and add 15 mL of 0.8% ammonium chloride solution to each tube. 
	NOTE: This step is to lyse the remaining red blood cells.</li>
	<li>Incubate on ice for 10 min.</li>
	<li>Wash the cells as before by topping up tubes with PBS and centrifuge for 5 min at 560 x g and 4 &#176;C with low brake. Aspirate the supernatant from both tubes.</li>
	<li>Resuspend each cell pellet in 25 mL of PBS and repeat to wash a second time.</li>
	<li>Resuspend each cell pellet in 6 mL of EGM-2 culture medium supplemented with 10% fetal bovine serum (FBS) and 1x antibiotic/antimycotic solution containing 10,000 U/mL penicillin, 10 mg/mL streptomycin and 25 &#181;g/mL amphotericin. 
	NOTE: EGM-2 culture medium consists of EBM-2 culture medium supplemented with 200 &#181;L of hydrocortisone, 2 mL of human fibroblast growth factor (hFGF-B), 500 &#181;L of vascular endothelial growth factor (VEGF), 500 &#181;L of recombinant analog of insulin-like growth factor (R3 IGF-1), 500 &#181;L of ascorbic acid, 500 &#181;L of human epidermal growth factor (hEGF), 500 &#181;L of gentamicin/amphotericin, and 500 &#181;L of heparin per 500 mL.</li>
	<li>Gently transfer 2 mL of the cell suspension to each well of the fibronectin-coated 6-well plate and gently rock until evenly coated. 
	&#8203;NOTE: The goal is to seed as many cells as possible to maximize the number of endothelial colonies that will form.</li>
</ol>
4. Cell culture
<ol>
	<li>Incubate the cells at 37 &#176;C, 5% CO2, and 100% humidity.</li>
	<li>On the next day, gently add 1 mL of fresh culture medium to each well. 
	NOTE: it is important to be gentle not to disturb cells that are loosely adherent to the fibronectin coating.</li>
	<li>On the next day, perform a half culture medium change by gently aspirating 1.5 mL of culture medium from each well and replacing it with 1.5 mL of fresh culture medium.</li>
	<li>On each of the next 5 days, gently perform a full culture medium change to each well (2 mL of fresh culture medium per well).</li>
	<li>After that, gently change the culture medium three times per week.</li>
	<li>Regularly visualize the cell cultures under low power (4x objective) light microscopy. 
	NOTE: Adherent blood outgrowth endothelial cell (BOEC) colonies will begin to appear in the wells 7-10 days after plating. Non-adherent cell types will be discarded with medium changes, and other adherent cell types will gradually dissipate as the BOEC colonies grow.</li>
	<li>Passage the BOECs into a fibronectin-coated T-75 flask when ~70%-80% confluent and continue changing culture medium three times per week. 
	NOTE: Seeding density can be controlled by passaging colonies into a T25 flask instead. Subsequent passages do not require fibronectin-coating, and culture medium changes can be reduced to two times per week.</li>
	<li>Cells from passages 1-3 may be used for experiments or transferred to a freezing medium and stored in liquid nitrogen for future use.</li>
</ol>

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The authors have nothing to disclose.

BOECs are a powerful tool that may be used in various scientific and therapeutic approaches7,8,16. BOECs have been used to analyze EC gene expression to elucidate the key factors responsible for the development of vascular diseases and cancer5,19,20,21. BOECs have also been used in therapeutic applications such as vascular regeneration and gene delivery22,23,24. Genetically modified BOECs have been known for their antiangiogenic tumor therapeutic ability25,26. Further, the angiogenic potential of BOECs makes them excellent candidates for cellular therapies aiming at vascularization and neovascularization17,27. Even though several studies describe how to obtain BOECs from human peripheral blood6,7,8,9, the variability of human cell sources due to underlying pathophysiological and genetic variation suggests a critical need to obtain BOECs from healthy xenogenic sources9,13,18.
A detailed protocol to efficiently derive BOECs from porcine peripheral blood mononuclear cells has been presented. Critical steps include processing blood samples as soon as possible and efficient harvesting of buffy coat layers after density gradient centrifugation to maximize the number of viable PBMCs going into the culture. The initial plating density and timely passaging of cells are important considerations for obtaining consistent isolation of these cell types. A starting blood volume of 100 mL provides an appropriate number of PBMCs for a single six-well plate. It should be noted that the maximum volume of blood that can be safely collected from a pig will vary with animal body weight.&#160;A suggested safe one-time maximum blood volume collection for pigs is 8 mL per kg of body weight. In addition, timely passaging of initial colonies from passage 0 to passage 1 is also essential. The cells cease to proliferate if seeding density is very low or if the cells reach confluency. As the cells become confluent, they start transforming into an elongated mesenchymal morphology and phenotype. Seeding density can also be controlled by passaging colonies into a T25 flask rather than a T75 flask. Careful consideration is required not to lose the adherent cell types during initial media change as it is very imperative to capture as many adherent cells as possible while discarding the non-adherent EPCs. This protocol produces outgrowth cell colonies in approximately 1 week.
The protocol is highly reproducible and reliable. Expanding endothelial cell colonies are achieved in an estimated 90%-95% of cultures. Failure is typically due to lack of formation of endothelial cell colonies, meaning if at least one endothelial cell colony is formed, they will nearly always proceed to expand in culture. Sources of failure include random variability, excessive shearing of cells during blood collection, contaminating microbes in the culture, and high loss of mononuclear cells during the processing steps. After an unsuccessful culture, a second culture will typically be successful, indicating that variability between animals is not a major factor. The protocol is even successful when collecting blood immediately after euthanasia, indicating the washing steps efficiently reduce the euthanasia agents to innocuous levels.
Irrespective of the heterogeneity in the circulating blood cell population, BOECs that are very distinct from EPCs were obtained. BOECs are derived from a small subset of adherent PBMCs that lack CD14 expression. This was confirmed by the absence of CD14 staining (Figure 3C,D). The clearance of monocytic EPCs from BOEC cultures in early passages may be explained either by cell death or non-adherence. The cobblestone morphology of BOECs (Figure 1E,F) suggests a pronounced endothelial phenotype. In addition, the uniform expression of surface marker CD31 (Figure 3A) implies a homogenous population of mature endothelial cells. Further, the capillary morphogenesis of BOECs both in serum-free (Figure 2A,B) and complete growth media (Figure&#160;2C,D) shows their inherent angiogenic capability6,8. Previous analysis of porcine BOECs generated using this protocol has shown these cells are additionally positive for von Willebrand factor expression and lectin binding28.
The abundance of PBMCs in circulating blood, from which BOECs emerge, supports the feasibility of a robust method to obtain a virtually limitless supply of therapeutic BOECs from animal sources. Once the technique is mastered, BOECs can be used for several applications, including vascular repair and regeneration, disease modeling, drug screening, and endothelial cell biology studies. Ongoing studies are needed to confirm the homogeneity of porcine BOECs and their compatibility as a therapeutic source in humans.
One limitation of this protocol is the risk of expanding endothelial cell colonies failing to generate. The failure rate is estimated to be 5%-10%, and in these instances, a second attempt is necessary. When assurance of success is important, twice the amount of blood can be collected, and the protocol can be followed in parallel to yield two independent cultures. Furthermore, our experience is limited to healthy Yorkshire/Landrace/Duroc cross domestic pigs (Sus domesticus), male and female, 40-80 kg, 3-6 months old. Success rates with other breeds, ages, and experimental conditions are unknown.
Another limitation of the protocol is that a definitive marker for BOEC phenotype does not exist. Here and in previous work28, the porcine BOECs generated from this protocol have been characterized for several BOEC markers, but the understanding of BOEC phenotype continues to evolve. BOECs are also characterized at passages 2-3 and phenotype at higher passages is unknown. An advantage of porcine BOECs compared to humans is the ready supply of pig blood, which allows for the generation of more frequent cultures and a robust supply of consistent, low passage cells.
One of the major limitations of using porcine cells in therapeutic approaches is the host immune response29. Several immunomodulation strategies are being investigated to mitigate the immunogenicity of xenogeneic cells for human transplantation, such as anti-inflammatory agents and CRISPR/Cas genetic manipulation30,31,32.&#160;More recent studies have proposed the expression of human complementary regulatory proteins (hCRPs)33 and the introduction of &#945;1,3- galactosyltransferase deleted (GTKO)32,34 pigs. These advancements hold promise for the therapeutic application of xenogeneic cells, including porcine BOECs.

The endothelium is a highly complex, dynamic structure and a vital component of the vascular wall. It lines the inner surface of blood vessels to provide a physical interface between circulating blood and surrounding tissues. This heterogeneous structure is known to perform various functions such as angiogenesis, inflammation, vasoregulation, and hemostasis1,2,3,4. Human umbilical vein endothelial cells are a widely studied cell type to assess the functionality of endothelial cells. However, the patient-specific batch variability, inconsistent phenotype, and minimum splitting efficiency suggest a need to determine a cell source that could improve upon all of these features5.
Obtaining a homogenous population of primary endothelial cells can be technically challenging, and primary endothelial cells do not possess high proliferative capacity6.&#160;Hence, to study vascular regeneration and evaluate pathophysiological processes, various groups have tried to obtain and assess different types of endothelial cells derived from peripheral blood, e.g., endothelial progenitor cells (EPCs) or blood outgrowth endothelial cells (BOECs)6,7,8,9. The spindle-shaped early EPCs originate from hematopoietic stem cells (HSCs) and have limited growth potency and limited angiogenic ability to produce mature endothelial cells. Furthermore, they closely resemble inflammatory monocytes. In addition, their capacity to further differentiate into functional, proliferating, mature endothelial cells is still debatable6,7,9,10.&#160;The continuous culture of peripheral blood mononuclear cells (PBMCs) can give rise to a secondary population of cells known as late-outgrowth EPCs, BOECs, or endothelial colony-forming cells (ECFCs)6,7,9,10. Medina et al. in 2018, acknowledged the limitations of EPCs, the ambiguity of their nomenclature, along with a general lack of concordance with many distinct cell types continuously grouped under EPCs11. In contrast, BOECs have become recognized for their role in vascular repair, health and disease, and cell therapy. Further study and therapeutic use of these cells will rely on protocols to consistently derive these cell types from circulating progenitor cells.
Primary cells such as BOECs can be used as a surrogate to obtain highly proliferative mature endothelial cells6. BOECs are phenotypically distinct from early EPCs and exhibit typical endothelial features such as cobblestone morphology and expression of adherens junctions and caveolae12. Gene profiling by Hebbel et al.13,14,15 found that BOECs or ECFCs are the true endothelial cells as they promote microvascular and large vessel formation. Thus, BOECs can be used as a tool to evaluate pathophysiological processes and genetic variation16. They are also considered an excellent cell source for cell therapy for vascular regeneration17. Hence, a standardized protocol to consistently derive these highly proliferative cells is essential.
While BOECs provide a powerful tool for studying human pathophysiological and genetic variation, a more homogenous source of BOECs may provide more robust and reliable experimental and therapeutic outcomes. Superior homogeneity can be achieved by using xenogeneic cell sources derived from genetically similar animals raised in similar conditions18. While xenogeneic cell sources are prone to eliciting a host immune response, immunomodulation strategies are being developed with the goal of generating immunocompatible animals and animal products, including cells. Pigs, in particular, are an abundant source of peripheral blood and are commonly used to study medical devices and other therapies due to anatomical and physiological similarities to humans. Hence, this study refines the protocol for the isolation and expansion of highly proliferative BOECs from porcine peripheral blood.The protocol detailed below is a straightforward and reliable method to obtain a large number of BOECs from a relatively small volume of blood. The cultures can be expanded through several passages to generate millions of cells from a single blood sample.

<table><tbody><tr><td>19 G needle</td><td>Covidien</td><td>1188818112</td><td></td></tr><tr><td>50 mL conical tubes</td><td>Corning</td><td>352098</td><td></td></tr><tr><td>6 well plate</td><td>BD Falcon</td><td>353046</td><td></td></tr><tr><td>60 mL syringes</td><td>Covidien</td><td>8881560125</td><td></td></tr><tr><td>Ammonium chloride solution (0.8%)</td><td>Stemcell Technologies</td><td>07850</td><td></td></tr><tr><td>Antibiotic/antimycotic solution (100x)</td><td>Gibco</td><td>15240-062</td><td></td></tr><tr><td>Centrifuge</td><td>Thermo Scientific</td><td>75-253-839</td><td></td></tr><tr><td>EGM-2 culture medium</td><td>Lonza Walkersville</td><td>CC-3162</td><td></td></tr><tr><td>Extension tube</td><td>Hanna Pharmaceutical Supply Co.</td><td>03382C6227</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Atlas Biologicals</td><td>F-0500-A</td><td></td></tr><tr><td>Ficoll-Paque 1077</td><td>Cytiva</td><td>17144003</td><td>Density gradient solution</td></tr><tr><td>Heparin sodium injection (1,000 units/mL)</td><td>Pfizer</td><td>00069-0058-01</td><td></td></tr><tr><td>Human plasma fibronectin</td><td>Gibco</td><td>33016-015</td><td></td></tr><tr><td>Ice</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Phosphate-buffered saline (PBS)</td><td>Gibco</td><td>10010-023</td><td></td></tr><tr><td>Pipette set</td><td>Eppendorf</td><td>2231300004</td><td></td></tr><tr><td>Sterile water</td><td>Gibco</td><td>15230-162</td><td></td></tr><tr><td>Thin pipette</td><td>Celltreat Scientific</td><td>229280</td><td></td></tr></tbody></table>

characterization of blood outgrowth endothelial cells (boec) from porcine peripheral blood

The endothelium is a dynamic integrated structure that plays an important role in many physiological functions such as angiogenesis, hemostasis, inflammation, and homeostasis. The endothelium also plays an important role in pathophysiologies such as atherosclerosis, hypertension, and diabetes. Endothelial cells form the inner lining of blood and lymphatic vessels and display heterogeneity in structure and function. Various groups have evaluated the functionality of endothelial cells derived from human peripheral blood with a focus on endothelial progenitor cells derived from hematopoietic stem cells or mature blood outgrowth endothelial cells (or endothelial colony-forming cells). These cells provide an autologous resource for therapeutics and disease modeling. Xenogeneic cells may provide an alternative source of therapeutics due to their availability and homogeneity achieved by using genetically similar animals raised in similar conditions. Hence, a robust protocol for the isolation and expansion of highly proliferative blood outgrowth endothelial cells from porcine peripheral blood has been presented. These cells can be used for numerous applications such as cardiovascular tissue engineering, cell therapy, disease modeling, drug screening, studying endothelial cell biology, and in vitro co-cultures to investigate inflammatory and coagulation responses in xenotransplantation.

Morphology of the cultured cells was observed from the start of the culture until BOEC colonies were observed (Figure 1). A smaller population of adherent cells started to attach to the culture dishes and grow, while non-adherent cells were removed with culture medium changes (Figure 1B). Colonies first appeared on day 6 as a collection of endothelial-like cells proliferating radially outward from a central point (Figure 1D). As the culture progressed, cell colonies became more dense and displayed a cobblestone morphology similar to mature endothelial cells (Figure 1F). The typical endothelial cobblestone morphology provides an easy preliminary identification of BOECs using a light microscope.
Endothelial cell colonies are typically ready for passaging at 10-14 days of culture. At this time, the 6-well plate will typically yield ~1 million cells with percent viability of 93%-98%. During the first passage, endothelial cells will expand to yield ~6-10 million cells in a T75 flask.
To assess the morphogenesis of BOECs, the basement membrane matrix (e.g., Matrigel) was plated on 15 well angiogenesis plates at 10 &#181;L/well and incubated at 37 &#176;C for 30 min to allow polymerization. Passage 2 cells were seeded on basement membrane matrix-coated plates and imaged at 14 h and 24 h time points. A density of approximately 3,500 cells per well was able to form a network and tube-like structures. Tube formation was noticed within 14 h for serum free medium (EGM-2 with supplementals except for FBS) and within 24 h for complete growth medium (EGM-2 with supplements including FBS). Adding FBS may have diluted the endothelial cell-specific growth factors (e.g., VEGF) and introduced other signaling factors resulting in the delay of the tube formation process. 2D phase-contrast microscopy was used to image tube formation in serum-free medium (Figure 2A,B) and complete growth medium (Figure 2C,D). Cells in both media conditions underwent morphological differentiation and rapidly organized into extensive networks of capillary tube-like structures. These structures were composed of organized cellular cords resembling in vivo capillary networks. Furthermore, micrographs at 20x magnification illustrate the complex multicellular organization of endothelial cells and their morphological differentiation in detail. There were no morphological differences observed between the media conditions. The extensive network of capillary tube-like structures strongly suggests the differentiation of cultured cells into mature and functional endothelial cells.
BOECs were further characterized by the expression of mature endothelial cell marker CD31 or platelet endothelial cell adhesion molecule-1 (PECAM1) (Figure 3A,B). BOECs showed uniform expression of CD31. Furthermore, flow cytometry analysis confirmed the absence of EPCs as the cells stained negative for monocyte marker CD14 compared to the positive control group of PBMCs (Figure 3C,D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63285/63285fig01.jpg" /> 
Figure 1: Phenotype light microscopy images. The light microscopy images of cultured BOECs observed at 10x magnification on (A) day 0, (B) day 2, (C) day 4, (D) day 6, (E) day 8, and (F) after the first passage. Scale bars: 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63285/63285fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63285/63285fig02.jpg" /> 
Figure 2: Morphological differentiation and organization of passage 2 BOECs. Morphological differentiation and organization of passage 2 BOECs into capillary tube-like structures was noticed in the basement membrane matrix within 14 h for serum-free medium and within 24 h for complete growth medium. (A) Serum-free medium at 4x, (B) Serum-free medium at 20x, (C) Complete growth medium at 4x, and (D) Complete growth medium at 20x. Scale bars: 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63285/63285fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63285/63285fig03.jpg" /> 
Figure 3: Characterization of BOECs with CD31 and CD14. (A) Passage 2-3 BOECs were stained for PECAM1 using anti-CD31-FITC antibody seen in green and (B) corresponding phase-contrast microscopy images. Cells were stained in suspension, and the suspension was imaged on a microscope slide. Scale bars: 200 &#181;m. (C) Flow cytometry analysis of BOECs with CD14-AF700 antibody compared to (D) positive control peripheral blood mononuclear cells (PBMCs). Positive CD14 staining shown in blue and unstained cells shown in red. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63285/63285fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Characterization of Blood Outgrowth Endothelial Cells BOEC from Porcine Peripheral Blood at JoVE.com

Characterization of Blood Outgrowth Endothelial Cells BOEC from Porcine Peripheral Blood

Characterization of Blood Outgrowth Endothelial Cells (BOEC) from Porcine Peripheral Blood

The endothelium is a dynamic integrated structure that plays an important role in many physiological functions such as angiogenesis, hemostasis, ...

characterization-blood-outgrowth-endothelial-cells-boec-from-porcine

Nanyang Technological University

Research

JoVE Journal

Bioengineering

1.6K Views.  Medical College of Wisconsin & Marquette University. Het protocol biedt zeer proliferatieve bloeduitgroei endotheelcellen die kunnen worden gebruikt voor verschillende weefselmanipulatie, celtherapie en ziektemodelleringsonderzoeken. Deze techniek levert consequent veel cellen op uit een enkele bloedafname. Om deze techniek succesvol te laten zijn, is het cruciaal om de cellen zo snel mogelijk in het kweekmedium en de incubatoromgeving te krijgen.Verdun eerst de heparineoplossing tot 100 eenheden per milliliter in steriele zoutoplossing....

Video: Characterization of Blood Outgrowth Endothelial Cells BOEC from Porcine Peripheral Blood

Autologous Endothelial Progenitor Cell-Seeding Technology and Biocompatibility Testing For Cardiovascular Devices in Large Animal Model

Implantable cardiovascular devices are manufactured from artificial materials (e.g. titanium (Ti), expanded polytetrafluoroethylene), which pose the risk of thromboemboli formation1,2,3. We have developed a method to line the inside surface of Ti tubes with autologous blood-derived human or porcine endothelial progenitor cells (EPCs)4. By implanting Ti tubes containing a confluent layer of porcine EPCs in the inferior vena cava (IVC) of pigs, we tested the improved biocompatibility of the cell-seeded surface in the prothrombotic environment of a large animal model and compared it to unmodified bare metal surfaces5,6,7 (Figure 1). This method can be used to endothelialize devices within minutes of implantation and test their antithrombotic function in vivo. 
Peripheral blood was obtained from 50 kg Yorkshire swine and its mononuclear cell fraction cultured to isolate EPCs4,8. Ti tubes (9.4 mm ID) were pre-cut into three 4.5 cm longitudinal sections and reassembled with heat-shrink tubing. A seeding device was built, which allows for slow rotation of the Ti tubes. 
We performed a laparotomy on the pigs and externalized the intestine and urinary bladder. Sharp and blunt dissection was used to skeletonize the IVC from its bifurcation distal to the right renal artery proximal. The Ti tubes were then filled with fluorescently-labeled autologous EPC suspension and rotated at 10 RPH x 30 min to achieve uniform cell-coating9. After administration of 100 USP/ kg heparin, both ends of the IVC and a lumbar vein were clamped. A 4 cm veinotomy was performed and the device inserted and filled with phosphate-buffered saline. As the veinotomy was closed with a 4-0 Prolene running suture, one clamp was removed to de-air the IVC. At the end of the procedure, the fascia was approximated with 0-PDS (polydioxanone suture), the subcutaneous space closed with 2-0 Vicryl and the skin stapled closed. 
After 3 - 21 days, pigs were euthanized, the device explanted en-block and fixed. The Ti tubes were disassembled and the inner surfaces imaged with a fluorescent microscope.
We found that the bare metal Ti tubes fully occluded whereas the EPC-seeded tubes remained patent. Further, we were able to demonstrate a confluent layer of EPCs on the inside blood-contacting surface.
Concluding, our technology can be used to endothelialize Ti tubes within minutes of implantation with autologous EPCs to prevent thrombosis of the device. Our surgical method allows for testing the improved biocompatibility of such modified devices with minimal blood loss and EPC-seeded surface disruption.

A method for seeding titanium blood-contacting biomaterials with autologous cells and testing biocompatibility is described. This method uses endothelial progenitor cells and titanium tubes, seeded within minutes of surgical implantation into porcine venae cavae. This technique is adaptable to many other implantable biomedical devices.

Implantable cardiovascular devices are manufactured from artificial materials (e.g. titanium (Ti), expanded polytetrafluoroethylene), which pose the risk ...

Parallel-plate Flow Chamber and Continuous Flow Circuit to Evaluate Endothelial Progenitor Cells under Laminar Flow Shear Stress

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well quantifiable fluid shear stresses1.
Our flow chamber design and flow circuit (Fig. 1) contains a transparent viewing region that enables testing of cell adhesion and imaging of cell morphology immediately before flow (Fig. 11A, B), at various time points during flow (Fig. 11C), and after flow (Fig. 11D). These experiments are illustrated with human umbilical cord blood-derived endothelial progenitor cells (EPCs) and porcine EPCs2,3.
This method is also applicable to other adherent cell types, e.g. smooth muscle cells (SMCs) or fibroblasts.
The chamber and all parts of the circuit are easily sterilized with steam autoclaving. In contrast to other chambers, e.g. microfluidic chambers, large numbers of cells (&gt; 1 million depending on cell size) can be recovered after the flow experiment under sterile conditions for cell culture or other experiments, e.g. DNA or RNA extraction, or immunohistochemistry (Fig. 11E), or scanning electron microscopy5. The shear stress can be adjusted by varying the flow rate of the perfusate, the fluid viscosity, or the channel height and width. The latter can reduce fluid volume or cell needs while ensuring that one-dimensional flow is maintained. It is not necessary to measure chamber height between experiments, since the chamber height does not depend on the use of gaskets, which greatly increases the ease of multiple experiments. Furthermore, the circuit design easily enables the collection of perfusate samples for analysis and/or quantification of metabolites secreted by cells under fluid shear stress exposure, e.g. nitric oxide (Fig. 12)6.

We are describing a method to subject adherent cells to laminar flow shear stress in a sterile continuous flow circuit. The cells' adhesion, morphology can be studied through the transparent chamber, samples obtained from the circuit for metabolite analysis and cells harvested after shear exposure for future experiments or culture.

The overall goal of this method is to describe a technique to subject adherent cells to laminar flow conditions and evaluate their response to well ...

Isolation of Murine Embryonic Hemogenic Endothelial Cells

The specification of hemogenic endothelial cells from embryonic vascular endothelium occurs during brief developmental periods within distinct tissues, and is necessary for the emergence of definitive HSPC from the murine extra embryonic yolk sac, placenta, umbilical vessels, and the embryonic aorta-gonad-mesonephros (AGM) region. The transient nature and small size of this cell population renders its reproducible isolation for careful quantification and experimental applications technically difficult. We have established a fluorescence-activated cell sorting (FACS)-based protocol for simultaneous isolation of hemogenic endothelial cells and HSPC during their peak generation times in the yolk sac and AGM. We demonstrate methods for dissection of yolk sac and AGM tissues from mouse embryos, and we present optimized tissue digestion and antibody conjugation conditions for maximal cell survival prior to identification and retrieval via FACS. Representative FACS analysis plots are shown that identify the hemogenic endothelial cell and HSPC phenotypes, and describe a methylcellulose-based assay for evaluating their blood forming potential on a clonal level.

Hematopoietic stem and progenitor cells (HSPC) derive from specialized (hemogenic) endothelial cells during development, yet little is known about the process by which some endothelial cells specify to become blood forming. We demonstrate a flow-cytometry based method allowing simultaneous isolation of hemogenic endothelial cells and HSPC from murine embryonic tissues.

The specification of hemogenic endothelial cells from embryonic vascular endothelium occurs during brief developmental periods within distinct tissues, ...

Developmental Biology

Isolation of Endothelial Progenitor Cells from Human Umbilical Cord Blood

The existence of endothelial progenitor cells (EPCs) in peripheral blood and its involvement in vasculogenesis was first reported by Ashara and colleagues1. Later, others documented the existence of similar types of EPCs originating from bone marrow2,3. More recently, Yoder and Ingram showed that EPCs derived from umbilical cord blood had a higher proliferative potential compared to ones isolated from adult peripheral blood4,5,6. Apart from being involved in postnatal vasculogenesis, EPCs have also shown promise as a cell source for creating tissue-engineered vascular and heart valve constructs7,8. Various isolation protocols exist, some of which involve the cell sorting of mononuclear cells (MNCs) derived from the sources mentioned earlier with the help of endothelial and hematopoietic markers, or culturing these MNCs with specialized endothelial growth medium, or a combination of these techniques9. Here, we present a protocol for the isolation and culture of EPCs using specialized endothelial medium supplemented with growth factors, without the use of immunosorting, followed by the characterization of the isolated cells using Western blotting and immunostaining.

The goal of this protocol is to isolate endothelial progenitor cells from umbilical cord blood. Some of the applications include using these cells as a biomarker for identifying patients with cardiovascular risk, treating ischemic diseases, and creating tissue-engineered vascular and heart valve constructs.

The existence of endothelial progenitor cells (EPCs) in peripheral blood and its involvement in vasculogenesis was first reported by Ashara and ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

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 ...

Biology

Phenotypic and Functional Characterization of Endothelial Colony Forming Cells Derived from Human Umbilical Cord Blood

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of circulating endothelial progenitor cells (EPCs) were first identified in adult human peripheral blood by Asahara et al. in 1997 2 bringing an infusion of new hypotheses and strategies for vascular regeneration and repair. EPCs are rare but normal components of circulating blood that home to sites of blood vessel formation or vascular remodeling, and facilitate either postnatal vasculogenesis, angiogenesis, or arteriogenesis largely via paracrine stimulation of existing vessel wall derived cells3. No specific marker to identify an EPC has been identified, and at present the state of the field is to understand that numerous cell types including proangiogenic hematopoietic stem and progenitor cells, circulating angiogenic cells, Tie2+ monocytes, myeloid progenitor cells, tumor associated macrophages, and M2 activated macrophages participate in stimulating the angiogenic process in a variety of preclinical animal model systems and in human subjects in numerous disease states4, 5. Endothelial colony forming cells (ECFCs) are rare circulating viable endothelial cells characterized by robust clonal proliferative potential, secondary and tertiary colony forming ability upon replating, and ability to form intrinsic in vivo vessels upon transplantation into immunodeficient mice6-8. While ECFCs have been successfully isolated from the peripheral blood of healthy adult subjects, umbilical cord blood (CB) of healthy newborn infants, and vessel wall of numerous human arterial and venous vessels 6-9, CB possesses the highest frequency of ECFCs7 that display the most robust clonal proliferative potential and form durable and functional blood vessels in vivo8, 10-13. While the derivation of ECFC from adult peripheral blood has been presented14, 15, here we describe the methodologies for the derivation, cloning, expansion, and in vitro as well as in vivo characterization of ECFCs from the human umbilical CB.

Endothelial colony forming cells (ECFCs) are circulating endothelial cells with robust clonal proliferative potential that display intrinsic in vivo vessel forming ability. Phenotypic and functional characterization of outgrowth endothelial cells derived from CB are important to identify and isolate bona fide ECFCs for potential clinical application in repairing damaged tissues.

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of ...

Assessment of the Anticoagulant and Anti-inflammatory Properties of Endothelial Cells Using 3D Cell Culture and Non-anticoagulated Whole Blood

In vivo, endothelial cells are crucial for the natural anticoagulation of circulating blood. Consequently, endothelial cell activation leads to blood coagulation. This phenomenon is observed in many clinical situations, like organ transplantation in the presence of pre-formed anti-donor antibodies, including xenotransplantation, as well as in ischemia/reperfusion injury. In order to reduce animal experimentation according to the 3R standards (reduction, replacement and refinement), in vitro models to study the effect of endothelial cell activation on blood coagulation would be highly desirable. However, common flatbed systems of endothelial cell culture provide a surface-to-volume ratio of 1 - 5 cm2 of endothelium per mL of blood, which is not sufficient for natural, endothelial-mediated anticoagulation. Culturing endothelial cells on microcarrier beads may increase the surface-to-volume ratio to 40 - 160 cm2/mL. This increased ratio is sufficient to ensure the &#34;natural&#34; anticoagulation of whole blood, so that the use of anticoagulants can be avoided. Here an in vitro microcarrier-based system is described to study the effects of genetic modification of porcine endothelial cells on coagulation of whole, non-anticoagulated human blood. In the described assay, primary porcine aortic endothelial cells, either wild type (WT) or transgenic for human CD46 and thrombomodulin, were grown on microcarrier beads and then exposed to freshly drawn non-anticoagulated human blood. This model allows for the measurement and quantification of cytokine release as well as activation markers of complement and coagulation in the blood plasma. In addition, imaging of activated endothelial cell and deposition of immunoglobulins, complement- and coagulation proteins on the endothelialized beads were performed by confocal microscopy. This assay can also be used to test drugs which are supposed to prevent endothelial cell activation and, thus, coagulation. On top of its potential to reduce the number of animals used for such investigations, the described assay is easy to perform and consistently reproducible.

We present an in vitro model which allows the study and analysis of coagulation in whole, non-anticoagulated blood. Anticoagulation in the system depends on the natural anticoagulation effect of healthy endothelial cells and endothelial cell activation will result in clotting.

In vivo, endothelial cells are crucial for the natural anticoagulation of circulating blood. Consequently, endothelial cell activation leads to blood ...

Medicine

Generation and Culture of Blood Outgrowth Endothelial Cells from Human Peripheral Blood

Historically, the limited availability of primary endothelial cells from patients with vascular disorders has hindered the study of the molecular mechanisms underlying endothelial dysfunction in these individuals. However, the recent identification of blood outgrowth endothelial cells (BOECs), generated from circulating endothelial progenitors in adult peripheral blood, may circumvent this limitation by offering an endothelial-like, primary cell surrogate for patient-derived endothelial cells. Beyond their value to understanding endothelial biology and disease modeling, BOECs have potential uses in endothelial cell transplantation therapies. They are also a suitable cellular substrate for the generation of induced pluripotent stem cells (iPSCs) via nuclear reprogramming, offering a number of advantages over other cell types. We describe a method for the reliable generation, culture and characterization of BOECs from adult peripheral blood for use in these and other applications. This approach (i) allows for the generation of patient-specific endothelial cells from a relatively small volume of adult peripheral blood and (ii) produces cells that are highly similar to primary endothelial cells in morphology, cell signaling and gene expression.

This protocol allows for the reliable generation and characterization of blood outgrowth endothelial cells (BOECs) from a small volume of adult peripheral blood. BOECs can be used as a surrogate for endothelial cells from patients with vascular disorders and as a substrate for the generation of induced pluripotent stem cells.

Historically, the limited availability of primary endothelial cells from patients with vascular disorders has hindered the study of the molecular ...

Isolation and Culture of Primary Aortic Endothelial Cells from Miniature Pigs

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a suitable organ source. Immune rejection and coagulation are two major hurdles for the success of xenotransplantation. Vascular endothelial cell (EC) injury and dysfunction are important for the development of the inflammation and coagulation responses in xenotransplantation. Thus, isolation of porcine aortic endothelial cells (pAECs) is necessary for investigating the immune rejection and coagulation responses. Here, we have developed a simple enzymatic approach for the isolation, characterization, and expansion of highly purified pAECs from miniature pigs. First, the miniature pig was anaesthetized with ketamine, and a length of aorta was excised. Second, the endothelial surface of aorta was exposed to 0.005% collagenase IV digestive solution for 15 min. Third, the endothelial surface of the aorta was scraped in only one direction with a cell scraper (&#60;10 times), and was not compressed during the process of scraping. Finally, the isolated pAECs of Day 3, and after passage 1 and passage 2, were identified by flow cytometry with an anti-CD31 antibody. The percentages of CD31-positive cells were 97.4% &#177; 1.2%, 94.4% &#177; 1.1%, and 92.4% &#177; 1.7% (mean &#177; SD), respectively. The concentration of Collagenase IV, the digestive time, the direction, and frequency and intensity of scraping are critical for decreasing fibroblast contamination and obtaining high-purity and a large number of ECs. In conclusion, our enzymatic method is a highly-effctive method for isolating ECs from the miniature pig aorta, and the cells can be expanded in vitro to investigate the immune and coagulation responses in xenotransplantation.

An effective enzymatic method for isolation of primary porcine aortic endothelial cells (pAECs) from miniature pigs is described. The isolated primary pAECs can be used to investigate the immune and coagulation response in xenotransplantation.

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a ...

Characterization of Blood Outgrowth Endothelial Cells (BOEC) from Porcine Peripheral Blood

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