Dr. Yoder is consultant to EndGenitor Technologies, Inc. and a member of the board of Rimedion Technologies, Inc.

Reagents and Solutions
EMG-2 media (Lonza, Cat. No. cc-3162 containing EBM-2 basal medium and EGM-2 SingleQuot kit Supplements, &amp; growth factors)
EBM-2 (Lonza, Cat. No. cc-3156) supplemented with the entire SingleQuot kit supplements &amp; growth factors (Lonza, Cat. No. cc-4176), 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin (10,000 U/ml)/streptomycin (10,000 &mu;g/ml)/amphotericin (25 &mu;g/ml). Store up to 1 month at 4 &deg;C. Recommended EGM-2 volumes to use for ECFC culture are 500 &mu;l/well for 24-well plates, 2 ml/well for 6-well plates, 5ml/25-cm2 flasks, and 10 ml/75-cm2 flasks, unless otherwise specified in the protocol.
Collagen I solution
Dilute 0.575 ml of glacial acetic acid (17.4N) in 495 ml of sterile distilled water (0.02 N final concentration). Sterile filter the dilute acetic acid with a 0.22-&mu;m vacuum filtration system. Add 25 mg rat tail collagen I to the dilute acetic acid to a final concentration of 50 &mu;g/ml. The amount of collagen added will vary depending on the collagen stock concentration. Store up to a month at 4 &deg;C.
Preparation of collagen I-coated tissue culture surfaces
Place 1 ml of collagen I solution in each well of a 6-well tissue culture plate (use 300 &mu;l/well for 24-well plates, 3ml/25-cm2 flasks, and 8 ml/75-cm2 flasks). Incubate 1 hour to overnight at 37 &deg;C. Remove the collagen I solution and wash surface two times, each time with PBS (use 500 &mu;l/well for 24-well plates, 2 ml/well for 6-well plates, 5ml/25-cm2 flasks, and 10 ml/75-cm2 flasks). Use plates immediately for cell cultures.
FACS staining buffer
Phosphate-buffered saline (PBS) supplemented with 2% (v/v) fetal bovine serum (FBS). Store up to 2 weeks at 4 &deg;C.
A. ECFC Outgrowth, Cloning and Expansion
<ol>
	<li>Collect CB with heparin (10 U heparin/ml blood) or EDTA as an anticoagulant and transport to the laboratory at room temperature and immediately (within 2 hours of infant's delivery) process the sample for mononuclear cell (MNC) isolation.</li>
	<li>Aliquot 15 ml of CB into each 50-ml conical centrifuge tube and add 20 ml of PBS to each tube of CB and pipette several times to mix. Draw up 15 ml Ficoll-Paque into a 20-ml syringe and attach a 20G needle or mixing cannula. Place the tip of the mixing cannula at the bottom of the tube of diluted blood and carefully underlay 15 ml Ficoll-Pague. Centrifuge the tubes 30 min at 740 x g, at room temperature, without the deceleration brake setting.</li>
	<li>Using a transfer pipette, remove the hazy layer of low-density MNCs located at the interface between the Ficoll-Paque and diluted plasma. Dispense the MNCs into a 50-ml conical tube containing 10 ml EGM-2 medium. Centrifuge the MNCs 10 min at 515 x g, at room temperature, with a high deceleration brake setting.</li>
	<li>Carefully aspirate and discard the supernatant. Wash the pelleted cells with EGM-2 twice (10 min at 515 x g) and re-suspend the MNCs in EGM-2 at 1.25 x 107 cells/ml. Prepare collagen I coated 6-well tissue culture plates by adding 1 ml rat-tail collagen type I (50 &mu;g/ml) per well and incubating the plate overnight. Pipette 4 ml of this MNC suspension into each well and place the cells in 37 &deg;C, 5% CO2 humidified incubator. A.1.5) After 24 hours (day 1), slowly remove the spent medium (do not disturb the loosely adherent cells) from the well with a pipette, slowly add 4 ml EGM-2 to the well, and return plate to the incubator. On day 2, refresh the medium by slowly removing the medium (do not disturb the loosely adherent cells) from the well with a pipette and adding 4 ml EGM-2 to each well. Repeat the medium change daily until day 7 and every other day thereafter until ECFC colonies appear for cloning. Typically colonies appear between day 5 and day 14 days7(Fig. 1).</li>
	<li>Visualize typical ECFC colonies with cobblestone morphology using inverted microscope (magnification 10x) and mark the colony boundaries with a marker on the underside of the well to indicate the position of each colony.</li>
	<li>On day 14, wash these primary colonies 2 times with PBS and harvest individual colonies using cloning cylinders and just enough TrypLE express to cover the cells inside the cylinders. After aspirating the final wash of PBS, place a sterile gel coated cloning cylinder around each colony and press firmly against the plate using sterile forceps. Add 2-3 drops of warm TrypLE express into each cloning cylinder and incubate for 3-5 min until cells within the cylinder begin to detach. Add approximately 250 &mu;l EGM-2 medium into the center of the cylinder and pipette up and down to generate single cell suspension. Transfer the single cell suspension from each cloning cylinder separately into an individual micro-centrifuge tube.</li>
	<li>Wash the area within the cylinder 2-3 times with approximately 250 &mu;l EGM-2 medium to collect all remaining cells and transfer the wash from each cylinder into the respective micro-centrifuge tube. Centrifuge the cells at high speed (&le;300 x g) on tabletop centrifuge for 5 min.</li>
	<li>Remove the supernatant and re-suspend the cell pellet in 1.5 ml fresh EGM-2 medium. Transfer the cell suspension (containing all the cells from individual colonies) from each tube into one well of a 24-well tissue culture plate pre-coated with 500 &mu;L rat-tail collagen I (50 &mu;g/ml).</li>
	<li>Place inside the incubator for expansion with media change performed every other day.</li>
	<li>When cells approach 80-90% confluency, remove the spent medium then wash the cells 2 times with PBS and add 500 &mu;l TrypLE express medium to each well of 24-well tissue culture plate. Place the plate inside the incubator for 3-5 min until cells begin to round up and detach. Add 1 ml of EGM-2 (with 10% defined FBS) to each well to collect the cells by washing and transfer them to a 15 ml tube. Wash the well with 1 ml EGM-2 medium to collect all remaining cells and transfer the wash from each well into respective 15 ml tube. Centrifuge the cells at 300 x g for 5 min.</li>
	<li>Remove the medium and re-suspend the cell pellet in 1 ml fresh EGM-2 medium (with 10% defined FBS). Obtain the viable cell count of an aliquot using a hemacytometer and trypan blue exclusion.</li>
	<li>Expand the cells by seeding about 5000 cells/cm2 onto a collagen I pre-coated tissue culture surfaces in EGM-2 media with media change every other day until cells approach 80-90% confluency before sub-culturing again.</li>
</ol>
B. In vitro Phenotypic Characterization of ECFCs: Endothelial Cell Surface Antigen Expression and Single Cell Assay for Clonal Proliferative Potential
<ol>
	<li>For endothelial cell surface antigen expression, detach cells with TrypLE express to collect cells by using methods described for ECFC cloning and expansion.</li>
	<li>Re-suspend cells in FACS staining buffer (10 X 106 cells/ 1ml FACS staining buffer) then add FcR blocking reagent (20 &mu;l/ 10 X 106 cells), mix gently and place on ice for 10 min.</li>
	<li>Aliquot 100 &mu;l of this cell suspension in micro-centrifuge tube for each endothelial surface antigen and isotype controls. Add appropriate amount of fluorochrome labeled human monoclonal antibodies that recognize endothelial antigens (CD31, CD144, CD146, and CD105) or hematopoietic antigens (CD45, and CD14) and leave on ice for 30 min protected from light. Caution: 1 X 106 cells are not required for each test. The authors typically prepare 0.5 to 1 X 105 cells/tube to obtain 2 X 104 analyzed events. Also, follow the manufacturer's recommendation for the amount of antibody to be used for each test. Some antibodies may require titrating to determine the optimal antibody concentration. All staining that the authors have performed are single color staining.</li>
	<li>After 30 min of incubation on ice, centrifuge each tube at high speed on a tabletop centrifuge for 5 min then remove the supernatant and re-suspend the cell pellet in FACS buffer.</li>
	<li>Analyze the samples on a flow cytometer to determine the percentage of cells that stain positively or negatively for each antigen. ECFCs stain uniformly positive for CD31, CD144, and CD146, but negative for CD45 and CD14 compared to the corresponding isotype controls.</li>
	<li>For single cell assay to examine the clonal proliferative potential, detach early passage (2-3) cells with TrypLE express to collect cells using methods similar to that was described for ECFC cloning and expansion.</li>
	<li>Infect these cells with enhanced green fluorescent protein (EGFP) expressing lentiviruses 16 and collect cells expressing EGFP by fluorescence cytometry. Caution: Working with lentivirus is considered a biohazard and all bio-safety precautions should be obeyed. </li>
	<li>Culture them as described for culturing ECFC in the section for ECFC expansion.</li>
	<li>Prepare Collagen I coated 96-well plates by adding 50 &mu;l rat-tail collagen type I (50 &mu;g/ml) per well and incubate the plate for overnight. Collect the EGFP+ ECFCs by using TrypLE express and resuspend them in EGM-2 medium with final concentration ~106 cells /ml.</li>
	<li>Use FACS Vantage (Becton Dickson) or other comparable sorter with low flow rate of 20 cells/second to sort one single EGFP+ ECFC per well of a 96-well plate and adjust the final volume to 200 &mu;l per well with EGM-2. Use an inverted fluorescence microscope to ensure that each well received only one cell. Alternatively, single cells can be sorted based on FSC and SSC and Sytox nuclear dye staining the colonies can be used for cell scoring and quantification.</li>
	<li>Incubate the plate over a period of two weeks with two media changes (200 &mu;l EGM-2 using multichannel pipetter) performed on day 5 and day 10. On day 14 of culture, wash each well with 100 &mu;l PBS before fixing the cells with 100 &mu;l of 4% paraformaldehyde for 30 min. For ECFCs that are not expressing EGFP, add Sytox reagent, a green fluorescent nuclear dye, following paraformaldehyde fixation and incubate at 4 &deg;C overnight.</li>
	<li>Use a fluorescence microscope to examine each well to quantitate the number of endothelial cells that expanded from a single cell cultured for 14 days. For quantitative analysis, score wells with 2 or more endothelial cells as positive (for proliferation) and examine them further for total endothelial cell count by visual counting with the fluorescence microscope.</li>
	<li>To display the obtained data, wells with an endothelial cell number of: 2 to 50, 51-2000, and 2001 or more, are considered as: endothelial cell clusters, low proliferative potential (LPP)-ECFCs, and high proliferative potential (HPP)-ECFCs respectively. ECFCs exhibit complete hierarchy of clonal proliferative potential by giving rise to endothelial clusters, LPPs, and HPPs when cultured at a single cell level for 14 days.7</li>
</ol>
C. In vivo Functional Characterization of ECFCs: In vivo Vessel Formation Assay to Examine the ECFC's Potential for Vasculogenesis
<ol>
	<li>Prepare cellularized gel implants by calculating the total volume (ml) of each of the following gel materials (with the final concentration in the parenthesis) needed to cast 1-ml gel (which will be later bisected to generate 2 implants): FBS (10%), EBM-2 10:1 (adjust the final volume to 1 ml), sodium bicarbonate (1.5 mg/ml), NaOH (adjust pH to 7.4), HEPES (25 mM), fibronectin (100 &mu;g/ml), and collagen I (1.5 mg/ml).</li>
	<li>After gel material calculation, detach cells with TrypLE express to collect cells using methods similar to that was described for ECFC cloning and expansion. Obtain a viable cell count of an aliquot using a hemacytometer and trypan blue. Transfer 2.4 million viable cells into a 50 ml-conical tube and pellet the cells by centrifugation at 515 x g, room temperature. Simultaneously, prepare gel matrix solution by adding calculated volumes of HEPES, sodium bicarbonate, EBM-2 10:1, FBS, fibronectin, and collagen to an ice-cold 50-ml conical tube (in the same order as they are listed). Mix thoroughly and adjust the pH to 7.4 with NaOH while keeping solution on ice.</li>
	<li>Discard the supernatant from the cells after centrifugation and re-suspend the pellet in 360 &mu;l warm EGM-2 and to it add pH adjusted 840 &mu;l of gel matrix solution, making the final volume of gel implant 1 ml. Mix the cells into gel matrix solution slowly until cells are thoroughly suspended in the gel solution.</li>
	<li>Transfer this 1 ml cellularized gel suspension to one well of a 12-well tissue culture plate and incubate for 20-30 min until the gel polymerizes. Gently cover the gel with 2 ml warm EGM-2 and incubate overnight.</li>
	<li>Immediately before the implantation, bisect the overnight incubated gel into two equal pieces using iris scissors and return the gel pieces to the same culture well containing EGM-2 medium.</li>
	<li>Use the animal surgical facility to sedate the animal (5-6 week old NOD/SCID mice) by administrating isoflurane anesthesia. Shave the lower part of abdomen and thoroughly clean the surgical site with alcohol pads and betadine or other antiseptic. Then isolate the area with drapes as per IACUC and institutional guidelines.</li>
	<li>Using sterile sharp iris scissors to make an approximately 5 mm incision in the lower quadrant of the abdomen, exposing the subcutaneous space between the skin and abdominal muscle. Perform blunt-dissection through the dermal layer from the abdominal muscle to create a 15-20 mm wide pocket leading superior into the upper abdominal quadrant. Repeat similar procedure to create similar open pocket in another side of the same mouse abdomen.</li>
	<li>Insert one half piece of gel into one side of abdominal pocket and another half piece of gel into another side of the abdominal pocket by lifting the dermal layer just caudal to the incision.</li>
	<li>After insertion of gels, close each incision with 2-3 stitches using 5-0 polypropylene suture on a cutting needle. Appropriately label the cage cards and perform post-surgical monitoring and analgesia as per the institutional and protocol requirements and allow 14 days for cells in these implanted gels to generate de novo vasculature.</li>
	<li>Finally, harvesting of gel implants is typically performed 14 days after implantation after euthanizing the mouse.</li>
	<li>Swab the abdominal area with alcohol pads and cut the abdominal skin caudal to the original incision line. Carefully, dissect out the implant by excising a flap of skin caudal to the probable location of the gel. Excise the implant by cutting circumferentially around the gel then place in zinc fixative (BD Biosciences, follow manufacturer's recommendation for optimal results) and allow them to fix for 1-2 hours at room temperature.</li>
	<li>Prepare the paraffin-embedded gels using standard histochemical protocols and prepare 5 &mu;m sections on glass slides to perform staining with hematoxylin and eosin, anti-human CD31 or anti-mouse CD31 to visualize the vasculature within the gel. ECFCs undergo de novo vasculogenesis to generate humanized vessels in cellularized gel implants inserted into immunodeficient mice for 14 days 4,8,11.</li>
</ol>
D. Representative Results
Using this ECFC derivation technique we have observed out-growth of primary colony formation as early as day 5 (Fig. 1). The out-growth ECFC colonies exhibited typical cobblestone appearance and gave rise to &gt;40 population doublings upon long term expansion after colony pickup by cloning. Expanded colonies expressed endothelial antigens, but did not express hematopoietic antigens (Fig. 2). Importantly, they displayed a complete hierarchy of clonal proliferative potential at a single cell level (Fig. 3). Moreover, ECFCs formed humanized blood vessels that are perfused with host RBCs when implanted into immunodeficient mice4, 6, 8, 11 (Fig. 4).
<img alt="Figure 1." src="/files/ftp_upload/3872/3872fig1.jpg" /> 
Figure 1. Isolation of mononuclear cells (MNCs) from cord blood and outgrowth of endothelial colony forming cells (ECFCs) from cultured MNCs. MNCs form buffy coat layer during Ficoll-Pague density gradient separation of cord blood cells. Isolation of buffy coat layer to culture MNCs on rat-tail collagen I coated plates results in outgrowth of ECFC colony in 5 to 14 days. The outgrowth ECFC colony (indicated by arrow heads) displayed cobblestone morphology7.
<img alt="Figure 2." src="/files/ftp_upload/3872/3872fig2.jpg" /> 
Figure 2. Representative in vitro phenotypic assessment of endothelial and hematopoietic cell surface antigen expression. Immunophenotyping of cord blood-derived ECFCs revealed that ECFCs expressed endothelial antigens CD31, CD34, CD144, CD146, Flt-1, Flk-1, Flt-4, and Nrp2 but did not express hematopoietic antigens CD45, CD14, CD11b, cKit, CXCR4 or AC1337, 8.
<img alt="Figure 3." src="/files/ftp_upload/3872/3872fig3.jpg" /> 
Figure 3. Representative in vitro quantitation of the clonogenic and proliferative potential of CB derived ECFCs. (A) cord blood-derived ECFCs display clonal proliferative potential with a hierarchy of colonies ranging from clusters of 2-50 cells up to colonies of &gt;2001. (B) Micrographs of hierarchy of colonies (colonies stained with, Sytox, a green fluorescent nuclear dye to take better quality pictures) obtained after cord blood-derived ECFCs were cultured at a single cell level for 14 days. Scale bar represents 100&mu;m7, 8.
<img alt="Figure 4." src="/files/ftp_upload/3872/3872fig4.jpg" /> 
Figure 4. Representative in vivo functional characterization of cord blood-derived ECFCs. A) H&amp;E staining of cord blood-derived ECFC containing cellularized gel implants indicated microvessel (filled with host RBCs) formation in collagen-fibronectin gel after 14 days of implantation. B) Anti-human CD31 staining (brown staining) further confirms the human origin of these vessels.

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No conflicts of interest declared.

Phenotypic and functional characterization of putative endothelial progenitor cells is important to identify the bona fide ECFCs that are capable of clonally and serially re-plating in culture and give rise to durable and functional implantable blood vessels in vivo. Human umbilical cord blood is enriched with ECFCs and the concentration of these circulating cells declines with aging or disease10. Recent studies suggest that ECFC may play important roles in vascular repair or regeneration in ...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
		</tr>
		<tr>
			<td>Heparin Sodium Injection, USP</td>
			<td>APP Pharmaceuticals</td>
			<td>504031</td>
		</tr>
		<tr>
			<td>Ficoll-Pague</td>
			<td>Amersham Biosciences</td>
			<td>17-1440-03</td>
		</tr>
		<tr>
			<td>Mixing cannula</td>
			<td>Maersk Medical</td>
			<td>500.11.012</td>
		</tr>
		<tr>
			<td>EGM-2</td>
			<td>Lonza</td>
			<td>CC-3162</td>
		</tr>
		<tr>
			<td>Defined FBS</td>
			<td>Hyclone</td>
			<td>SH30070.03</td>
		</tr>
		<tr>
			<td>TrypLE express</td>
			<td>Gibco</td>
			<td>12605</td>
		</tr>
		<tr>
			<td>Rat type I collagen</td>
			<td>BD Biosciences</td>
			<td>354236</td>
		</tr>
		<tr>
			<td>Matrigel</td>
			<td>BD Biosciences</td>
			<td>356234</td>
		</tr>
		<tr>
			<td>FcR Block</td>
			<td>Miltenyi Biotech</td>
			<td>130-059-901</td>
		</tr>
		<tr>
			<td>hCD31, FITC conjugated</td>
			<td>BD Pharmingen</td>
			<td>555445</td>
		</tr>
		<tr>
			<td>hCD45, FITC conjugated</td>
			<td>BD Pharmingen</td>
			<td>555482</td>
		</tr>
		<tr>
			<td>hCD14, FITC conjugated</td>
			<td>BD Pharmingen</td>
			<td>555397</td>
		</tr>
		<tr>
			<td>hCD144, PE conjugated</td>
			<td>eBioscience</td>
			<td>12-1449-80</td>
		</tr>
		<tr>
			<td>hCD146, PE conjugated</td>
			<td>BD Pharmingen</td>
			<td>550315</td>
		</tr>
		<tr>
			<td>hCD105, PE conjugated</td>
			<td>Invitrogen</td>
			<td>MHCD10504</td>
		</tr>
		<tr>
			<td>Ms IgG1,k antibody, FITC conjugated</td>
			<td>BD Pharmingen</td>
			<td>555748</td>
		</tr>
		<tr>
			<td>Ms IgG1,k antibody, PE conjugated</td>
			<td>BD Pharmingen</td>
			<td>559320</td>
		</tr>
		<tr>
			<td>Ms IgG2a,k antibody, FITC conjugated</td>
			<td>BD Pharmingen</td>
			<td>555573</td>
		</tr>
		<tr>
			<td>Anti-human CD31</td>
			<td>Dako</td>
			<td>clone JC70/A</td>
		</tr>
		<tr>
			<td>Anti-mouse CD31</td>
			<td>BD Pharmingen</td>
			<td>553370</td>
		</tr>
		<tr>
			<td>0.22-&mu;m vacuum filtration system</td>
			<td>Millipore</td>
			<td>SCGPU05RE</td>
		</tr>
		<tr>
			<td>Glacial acetic acid, 17.4N</td>
			<td>Fisher</td>
			<td>A38-500</td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic</td>
			<td>Invitrogen</td>
			<td>15240-062</td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Hyclone</td>
			<td>SH30070.03</td>
		</tr>
		<tr>
			<td>IHC Zinc Fixative</td>
			<td>BD Biosciences</td>
			<td>550523</td>
		</tr>
		<tr>
			<td>Sytox green reagent</td>
			<td>Invitrogen</td>
			<td>S33025</td>
		</tr>
		<tr>
			<td>Cloning cylinders, sterile</td>
			<td>Fisher Scientific</td>
			<td>07-907-10</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Phenotypic and Functional Characterization of Endothelial Colony Forming Cells Derived from Human Umbilical Cord Blood at JoVE.com

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

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 ...

phenotypic-functional-characterization-endothelial-colony-forming

Research

JoVE Journal

Biology

16.2K Views.  Indiana University School of Medicine. Эндотелиальных клеток, образующих колонии (ECFCs) циркулируют эндотелиальных клетках с надежными клонального пролиферативный потенциал, которые отображают внутренние В естественных условиях Судно формирование способности. Фенотипические и функцион?...

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

Isolation of Human Umbilical Vein Endothelial Cells (HUVEC)

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

Isolation of Human Umbilical Vein Endothelial Cells HUVEC

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

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

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

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

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

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

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

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

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

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

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

Isolation and Enrichment of Rat Mesenchymal Stem Cells (MSCs) and Separation of Single-colony Derived MSCs

MSCs are a population of adult stem cells that is a promising source for therapeutic applications. These cells can be isolated from the bone marrow and can be easily separated from the hematopoietic stem cells (HSCs) due to their plastic adherence. This protocol describes how to isolate MSCs from rat femurs and tibias. The isolated cells were further enriched against two MSCs surface markers CD54 and CD90 by magnetic cell sorting. Expression of surface markers CD54 and CD90 were then confirmed by flow cytometry analysis. HSC marker CD45 was also included to check if the sorted MSCs were depleted of HSCs. MSCs are naturally quite heterogeneous. There are subpopulations of cells that have different shapes, proliferation and differentiation abilities. These subpopulations all express the known MSCs markers and no unique marker has yet been identified for the different subpopulations. Therefore, an alternative approach to separate out the different subpopulations is using cloning cylinders to separate out single-colony derived cells. The cells derived from the single-colonies can then be cultured and evaluated separately.

Isolation and Enrichment of Rat Mesenchymal Stem Cells MSCs and Separation of Single-colony Derived MSCs

Rat MSCs were isolated from femurs and tibias and then enriched by magnetic cell sorting. Sorted cells were confirmed for the expression of surface markers by flow cytometry. These cells were also cultured at clonal density to form single colonies and then these colonies were separated by cloning cylinders.

MSCs are a population of adult stem cells that is a promising source for therapeutic applications. These cells can be isolated from the bone marrow and ...

Isolation of Human Umbilical Arterial Smooth Muscle Cells (HUASMC)

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded and their collection does not imply any added risk to the newborn or mother s health. Moreover no ethical concerns are raised. The UC is composed by one vein and two arteries from which both endothelial cells (ECs) 1 and smooth muscle cells (SMCs) 2, two of the main cellular components of blood vessels, can be isolated. In this project the SMCs were obtained after enzymatic treatment of the UC arteries accordingly the experimental procedure previously described by Jaffe et al 3. After cell isolation they were kept in t-flash with DMEM-F12 supplemented with 5% of fetal bovine serum and were cultured for several passages. Cells maintained their morphological and other phenotypic characteristics in the different generations. The aim of this study was to isolate smooth muscle cells in order to use them as models for future assays with constrictor drugs, isolate and structurally characterize L-type calcium channels, to study cellular and molecular aspects of the vascular function 4 and to use them in tissue engineering.

Isolation of Human Umbilical Arterial Smooth Muscle Cells HUASMC

The umbilical cords are used to isolate smooth muscle cells by different ways. In this work we used the enzymatic treatment to isolated smooth muscle cells.

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded ...

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

A Quantitative Assay for Insulin-expressing Colony-forming Progenitors

The field of pancreatic stem and progenitor cell biology has been hampered by a lack of in vitro functional and quantitative assays that allow for the analysis of the single cell. Analyses of single progenitors are of critical importance because they provide definitive ways to unequivocally demonstrate the lineage potential of individual progenitors. Although methods have been devised to generate "pancreatospheres" in suspension culture from single cells, several limitations exist. First, it is time-consuming to perform single cell deposition for a large number of cells, which in turn commands large volumes of culture media and space. Second, numeration of the resulting pancreatospheres is labor-intensive, especially when the frequency of the pancreatosphere-initiating progenitors is low. Third, the pancreatosphere assay is not an efficient method to allow both the proliferation and differentiation of pancreatic progenitors in the same culture well, restricting the usefulness of the assay.
To overcome these limitations, a semi-solid media based colony assay for pancreatic progenitors has been developed and is presented in this report. This method takes advantage of an existing concept from the hematopoietic colony assay, in which methylcellulose is used to provide viscosity to the media, allowing the progenitor cells to stay in three-dimensional space as they undergo proliferation as well as differentiation. To enrich insulin-expressing colony-forming progenitors from a heterogeneous population, we utilized cells that express neurogenin (Ngn) 3, a pancreatic endocrine progenitor cell marker. Murine embryonic stem (ES) cell-derived Ngn3 expressing cells tagged with the enhanced green fluorescent protein reporter were sorted and as many as 25,000 cells per well were plated into low-attachment 24-well culture dishes. Each well contained 500 &mu;L of semi-solid media with the following major components: methylcellulose, Matrigel, nicotinamide, exendin-4, activin &beta;B, and conditioned media collected from murine ES cell-derived pancreatic-like cells. After 8 to 12 days of culture, insulin-expressing colonies with distinctive morphology were formed and could be further analyzed for pancreatic gene expression using quantitative RT-PCR and immunoflourescent staining to determine the lineage composition of each colony.
In summary, our colony assay allows easy detection and quantification of functional progenitors within a heterogeneous population of cells. In addition, the semi-solid media format allows uniform presentation of extracellular matrix components and growth factors to cells, enabling progenitors to proliferate and differentiate in vitro. This colony assay provides unique opportunities for mechanistic studies of pancreatic progenitor cells at the single cell level.

A three-dimensional clonogenic assay that allows pancreatic-like progenitors to differentiate into insulin-expressing colonies is described. This method takes advantage of semi-solid media containing methylcellulose, Matrigel and growth factors, in which single progenitors proliferate and differentiate in vitro, permitting quantification of the number of functional progenitors in a population.

The field of pancreatic stem and progenitor cell biology has been hampered by a lack of in vitro functional and quantitative assays that allow for the ...

Manual Isolation of Adipose-derived Stem Cells from Human Lipoaspirates

In 2001, researchers at the University of California, Los Angeles, described the isolation of a new population of adult stem cells from liposuctioned adipose tissue that they initially termed Processed Lipoaspirate Cells or PLA cells. Since then, these stem cells have been renamed as Adipose-derived Stem Cells or ASCs and have gone on to become one of the most popular adult stem cells populations in the fields of stem cell research and regenerative medicine. Thousands of articles now describe the use of ASCs in a variety of regenerative animal models, including bone regeneration, peripheral nerve repair and cardiovascular engineering. Recent articles have begun to describe the myriad of uses for ASCs in the clinic. The protocol shown in this article outlines the basic procedure for manually and enzymatically isolating ASCs from large amounts of lipoaspirates obtained from cosmetic procedures. This protocol can easily be scaled up or down to accommodate the volume of lipoaspirate and can be adapted to isolate ASCs from fat tissue obtained through abdominoplasties and other similar procedures.

In 2001, researchers at UCLA described the isolation of a population of adult stem cells, termed Adipose-derived Stem Cells or ASCs, from adipose tissue. This article outlines the isolation of ASCs from lipoaspirates using a manual, enzymatic digestion protocol using collagenase.

In 2001, researchers at the University of  California, Los Angeles, described the isolation of a new population of adult  stem cells from liposuctioned ...

Isolation of Blood-vessel-derived Multipotent Precursors from Human Skeletal Muscle

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective isolation in culture. Recently, using fluorescence-activated cell sorting (FACS), we and other researchers prospectively identified and purified three subpopulations of multipotent precursor cells associated with the vasculature of human skeletal muscle. These three cell populations: myogenic endothelial cells (MECs), pericytes (PCs), and adventitial cells (ACs), are localized respectively to the three structural layers of blood vessels: intima, media, and adventitia. All of these human blood-vessel-derived stem cell (hBVSC) populations not only express classic MSC markers but also possess mesodermal developmental potentials similar to typical MSCs. Previously, MECs, PCs, and ACs have been isolated through distinct protocols and subsequently characterized in separate studies. The current isolation protocol, through modifications to the isolation process and adjustments in the selective cell surface markers, allows us to simultaneously purify all three hBVSC subpopulations by FACS from a single human muscle biopsy. This new method will not only streamline the isolation of multiple BVSC subpopulations but also facilitate future clinical applications of hBVSCs for distinct therapeutic purposes.

Blood vessels within human skeletal muscle harbor several multi-lineage precursor populations that are ideal for regenerative applications. This isolation method allows simultaneous purification of three multipotent precursor cell populations respectively from three structural layers of blood vessels: myogenic endothelial cells from intima, pericytes from media, and adventitial cells from adventitia.

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective ...

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on changes in intracellular calcium ([Ca2+]i) and nitric oxide (NO). In order to understand how [Ca2+]i, NO and downstream molecules are handled by a blood vessel in response to vasoconstrictors and vasodilators, we developed a novel technique that applies calcium-labeling (or NO-labeling) dyes with two photon microscopy to measure calcium handling (or NO production) in isolated blood vessels. Described here is a detailed step-by-step procedure that demonstrates how to isolate an aorta from a rat, label calcium or NO within the endothelial or smooth muscle cells, and image calcium transients (or NO production) using a two photon microscope following physiological or pharmacological stimuli. The benefits of using the method are multi-fold: 1) it is possible to simultaneously measure calcium transients in both endothelial cells and smooth muscle cells in response to different stimuli; 2) it allows one to image endothelial cells and smooth muscle cells in their native setting; 3) this method is very sensitive to intracellular calcium or NO changes and generates high resolution images for precise measurements; and 4) described approach can be applied to the measurement of other molecules, such as reactive oxygen species. In summary, application of two photon laser emission microscopy to monitor calcium transients and NO production in the endothelial and smooth muscle cells of an isolated blood vessel has provided high quality quantitative data and promoted our understanding of the mechanisms regulating vascular function.

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Vascular cell functiondepends on activity of intracellular messengers. Described here is an ex vivo two photon imaging method that allows the measurement of intracellular calcium and nitric oxide levels in response to physiological and pharmacological stimuli in individual endothelial and smooth muscle cells of an isolated aorta.

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on ...