This material is based upon work supported by the National Science Foundation under Grant No. CMMI-1452943 and by the University of Arkansas Honors College. We would also like to acknowledge the Arkansas Cord Blood Bank for providing us with cord blood units.

This research was carried out with the approval of the University of Arkansas Institutional Review Board (Approval number 16-04-722). Umbilical cord blood units were collected in citrate phosphate dextrose (CPD) solution at the Arkansas Cord Blood Bank, and units that did not meet the requirement for storage were donated for research. Cord blood units were couriered to the lab within 24 h of collection at ambient temperatures.
1. Isolation of Endothelial Progenitor Cells from Cord Blood
<ol>
	<li>Preparation of reagents.
	<ol>
		<li>Prepare EGM by adding endothelial basal medium (EBM) to 10% fetal bovine serum (FBS) supplemented with 20 ng/mL of insulin-like growth factor (IGF), 1 &#181;g/mL ascorbic acid, 5 ng/mL recombinant human epidermal growth factor (rhEGF), 22.5 &#181;g/mL heparin, and 0.5 ng/mL vascular endothelial growth factor (VEGF), along with 10 ng/mL of recombinant human Fibroblast growth factor B (rhFGF-B), 0.2 &#181;g/mL hydrocortisone, and 2% penicillin-streptomycin-glutamine. Sterilize the culture medium using a 0.2 &#181;m polyethersulfone (PES) membrane vacuum filter.</li>
		<li>Prepare rat tail I collagen solution at a concentration of 50 &#181;g/mL using 0.02 N acetic acid. Sterilize this solution using a 0.2 &#181;m syringe filter.</li>
		<li>Pre-coat 6-well plates with 2 mL of prepared rat tail I collagen solution for each well. Place them in an incubator maintained at 5% CO2 and 37 &#176;C for at least 1 h before seeding.</li>
		<li>Dilute approximately 25 mL of cord blood with Hanks Balanced Salt Solution (HBSS) at a 1:1 concentration.</li>
		<li>Prepare radio-immuno precipitation assay (RIPA) Buffer using 150 mM sodium chloride, 1% Triton X-100, 0.5% sodium deoxycholate, 1 mM &#946;-glycerophosphate, 0.1% SDS, and 50 mM Tris (pH 8.0).</li>
	</ol>
	</li>
	<li>Isolation of endothelial progenitor cells.
	<ol>
		<li>Warm all prepared reagents in a water bath maintained at 37 &#176;C prior to isolation.</li>
		<li>Add 20 mL of density gradient medium reagent to a 50 mL centrifuge tube.</li>
		<li>Carefully layer 20 mL of diluted cord blood in the centrifuge tube containing density gradient medium without breaking this layer by aiming the pipette towards the side walls of the tube.</li>
		<li>Separate the components of blood based on their densities (Figure 1) by centrifuging at 800 x g for 30 min at room temperature, with the brakes of the centrifuge rotor off. Do not disturb the different layers.</li>
		<li>Remove the plasma layer by carefully pipetting it out of the tube without disturbing other cell layers until the pipette tip is able to reach the MNC layer (see Figure 1). Discard the removed plasma. 
		NOTE: While using pipette tips to remove the plasma, leave a small layer of plasma above the MNC layer (Figure 1) to reduce disturbances.</li>
		<li>Collect the buffy coat that contains the MNCs using a syringe fitted to an 18-gauge needle and transfer it to a fresh centrifuge tube.</li>
		<li>Add an equal volume of EBM to the collected MNCs. Centrifuge these tubes at 500 x g for 10 min at 4 &#176;C. Discard the supernatant. 
		NOTE: The cell pellet will appear red because of the presence of red blood cells/erythrocytes.</li>
		<li>Add 5 mL of ammonium chloride solution to lyse the red blood cells. Incubate this tube on ice for 5 - 10 min, with occasional shaking. 
		NOTE: Caution should be taken during this step to not exceed the time duration, as it can be harmful to the MNCs.</li>
		<li>Centrifuge the tubes at 500 x g for 10 min at 4 &#176;C and discard the supernatant. If erythrocytes persist, repeat steps 1.2.8 and 1.2.9 until no red color is observed in the pellet.</li>
		<li>Resuspend the pellet in prepared EGM and use the hemocytometer to count the MNCs.</li>
		<li>Aspirate the rat tail I collagen-coated 6-well plate (from step 1.1.3) and wash the wells three times with 1x phosphate-buffered saline (PBS).</li>
		<li>Seed the MNC pellet resuspended in EGM to the collagen plate at a concentration of 1 x 107 MNCs in each well. Place it in the 37 &#176;C, 5% CO2 incubator.</li>
		<li>After 24 h, aspirate the medium and wash the wells once with EGM. Add 3 mL of EGM medium to each well and place it back in the 37 &#176;C, 5% CO2 incubator.</li>
		<li>Replenish the plate with fresh EGM medium every day for 7 days. After 7 days, change the EGM medium every other day.</li>
		<li>Using a bright-field microscope, take images of the 6-well plate every day to track the progression of endothelial cell colonies. Mark the plate where the colonies arise to keep track of their growth. 
		NOTE: It usually takes between 5 and 9 days for the colonies to appear in culture.</li>
	</ol>
	</li>
	<li>Expansion of endothelial progenitor cells.
	<ol>
		<li>Coat the T-25 culture flask with rat tail I collagen (50 &#181;g/mL) and place it in the 37 &#176;C, 5% CO2 incubator for at least 60 min. Aspirate and wash the T-25 cell culture flask three times with 1x PBS prior to seeding the cells.</li>
		<li>Trypsinize the endothelial cell colonies when they reach about 3 mm in size using 150 &#181;L of 0.05% Trypsin-EDTA solution in each well. Place the 6-well plate back into the 37 &#176;C, 5% CO2 incubator for 2 - 3 min.</li>
		<li>Gently tap the 6-well plate to dislodge the attached cells. Immediately add 2 mL of EGM and recover the cells in a 15 mL centrifuge tube.</li>
		<li>Count the cells using a hemocytometer and calculate the initial seeding density.</li>
		<li>Spin the 15 mL centrifuge tubes at 400 x g for 5 min. Resuspend the cell pellet in EGM and seed the T-25 flask with ~500,000 cells. Mark this flask as Passage 1 and place the T-25 flask back in the 37 &#176;C, 5% CO2 incubator. 
		NOTE: Cells can be plated in EBM and DMEM to compare the growth with EGM.</li>
		<li>For further passages, when the flask reaches 90% confluence, follow steps 1.3.1 - 1.3.5 and reseed the cells at 104 cells/cm2 on T-75 or T-175 cell culture flasks. Proceed to the characterization of the isolated cells (steps 2 and 3).</li>
	</ol>
	</li>
</ol>
2. Characterization of Isolated Cells Using Western Blotting
<ol>
	<li>Add 50 - 100 &#181;L of lysis buffer at every passage when following the expansion steps to characterize the isolated cells. Collect proteins from approximately 500,000 or more cells.</li>
	<li>Pipette up and down vigorously to rupture the cells and release the proteins. Collect and transfer the buffer to a microcentrifuge tube.</li>
	<li>Spin the microcentrifuge tube at 500 x g and carefully transfer the supernatant to a new microcentrifuge tube. Label and store the tubes at -80 &#176;C for long-term storage.</li>
	<li>Quantify the amount of proteins in each tube using either Bradford&#39;s or BCA assay27,28,29.</li>
	<li>Carry out Western blotting using standard procedures27,28,29. Analyze 5 &#181;g of total protein for the expression of CD31, CD34, VEGFR2, and &#945;-SMA. Use &#945;-tubulin for normalizing the data.</li>
</ol>
3. Indirect Immunofluorescence
<ol>
	<li>Sonicate 18-mm glass coverslips in 50% ethanol and dry them. Leave the clean coverslips in a 12-well plate overnight in the biosafety hood with a UV light on to sterilize them.</li>
	<li>Coat the coverslips with 50 &#181;g/mL of rat tail I collagen and transfer the plate to a 37 &#176;C, 5% CO2 incubator for at least 60 min.</li>
	<li>Aspirate the collagen and add 1 mL of 1x PBS to the 12-well plate to wash the coverslips. Aspirate the 1x PBS and repeat twice.</li>
	<li>Seed 250,000 cultured EPCs in each well and place them back in the 37 &#176;C, 5% CO2 incubator for at least 3 days before carrying out immunostaining.</li>
	<li>Wash the plate 3 times with 1x PBS. Add 1 mL of ice-cold methanol to each well and incubate at room temperature for 15 min to fix the cells.</li>
	<li>Carry out immunostaining using a standard protocol27,28,29. Use the antibodies at their appropriate dilutions (e.g., CD31 (1:20), CD34 (1:50), &#945;-SMA (1:100), and VEGFR2 (1:50)).</li>
	<li>Take images of the immunostained coverslips using an epifluorescence microscope.</li>
</ol>

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The authors have nothing to disclose.

As mentioned earlier, adherent EPCs possess a cobblestone morphology. Our isolated MNCs progressed from a spindle-shaped cell colony (Figure 2A-2D) in the early stages to a cobblestone colony (Figure 2E-2F) over a period of ten days in culture. EPCs have been labeled differently by different research groups, namely as late endothelial progenitor cells10, endothelial colony forming cells5...

Several investigators have studied the characteristics and potential of human EPCs5,10,11,12,13. EPCs can be described as circulating cells that have the ability to adhere to endothelial tissue in sites of hypoxia, ischemia, injury, or tumor formation and contribute to the formation of new vascular structures4,14. Their observed involvement in neovascularization, in the form of postnatal vasculogenesis, has led to an understanding of the pathophysiology of these cells and their use in therapeutic applications4,15,16. The number of EPCs in an individual has been shown to be correlated with cardiovascular pathology9,15,16,17,18,19,20. Other studies have also differentiated EPCs into a valve fibroblast-like phenotype and proposed that these cells could be used for tissue-engineering heart valves7,21.
The particular cell surface molecules needed to isolate EPCs have not been clearly identified due to discrepancies between investigations4. The adhesion of MNCs to a certain matrix, with exposure to a variety of culture conditions, has been performed by several groups1,17,22,23, suggesting that putative EPCs may display different phenotypic properties. These properties include a lack of phagocytotic ability, tube formation in Matrigel, and the uptake of Dil-acetylated low-density lipoproteins. The high clonogenic and proliferative potential are two properties with which EPCs can be hierarchized5. EPCs can also form in vitro tubules when cocultured with human fetal lung fibroblasts4. These cells are known to express endothelial cell surface markers and to share some of the hematopoietic markers13,24,25. The positively expressed markers that are widely accepted for phenotyping EPCs are CD31, CD34, vascular endothelial growth factor receptor 2 (VEGFR2), von Willebrand Factor (vWF), CD133, c-Kit, and vascular endothelial cadherin (VE-cadherin)4,18. Cells that co-express CD90, CD45, CD14, CD115, or alpha-smooth muscle actin (&#945;-SMA) are not considered to be EPCs because of their limited proliferative potential, ability to phagocytose bacteria, and inability to form de novo human vessels in vivo4,7. This article outlines a modified protocol for the isolation of endothelial progenitor cells from human umbilical cord blood without the need for any cell sorting protocols. For this article, we used CD31, CD34, and VEGFR2 as the positive markers, with &#945;-SMA as the negative indicator.
In this article, we propose a method of isolating and culturing endothelial progenitor cells from umbilical cord blood without cell sorting using specialized endothelial growth medium supplemented with growth factors (EGM). This EGM contains vascular endothelial growth factor (VEGF) and fibroblast growth factor (FGF), which enhance the survival, proliferation, and migration of endothelial cells26. It also includes ascorbic acid, which is responsible for maintaining the cobblestone morphology of cells; insulin-like growth factor-1 (IGF-1), which provides angiogenic and migratory function; and heparin, which causes improved long-term stability of growth factors in the medium26. Other growth factors added to the endothelial cell culture medium includes supplementation with epidermal growth factor (EGF), which helps in stimulating cell proliferation and differentiation, and hydrocortisone, which sensitizes the cells to EGF26. We show that the use of this specific growth medium yields higher numbers of EPCs compared to endothelial basal medium (EBM) or Dulbecco&#39;s Modified Eagle Medium (DMEM).

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		<tr height="20">
			<td height="20" style="width:204px;height:20px;">A) For isolation and culturing</td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td style="width:807px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">EGM-2 BulletKit</td>
			<td style="width:127px;">Lonza</td>
			<td>CC-3162</td>
			<td>This product comes with all the growth factors needed to make the Endothelial Growth Medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Fetal Bovine Serum</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">26140079</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Pencillin-Streptomycin-Glutamine (100X)</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">10378016</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">Ficoll-Paque</td>
			<td style="width:127px;">GE Heatlhcare</td>
			<td style="width:119px;">17-1440-02</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Hank&#39;s Balanced Salt Solution</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">14170-112</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Ammonium Chloride</td>
			<td style="width:127px;">Stem Cell Technologies</td>
			<td style="width:119px;">7850</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">1X Phosphate Buffer Saline</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">14190250</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">Rat Tail I Collagen</td>
			<td style="width:127px;">Corning</td>
			<td style="width:119px;">354236</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">Glacial Acetic Acid</td>
			<td style="width:127px;">Amresco</td>
			<td style="width:119px;">0714-500ML</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">0.05% Trypsin-EDTA</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">25300054</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">HEPES buffer</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">15630080</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Dulbecco&#39;s Modified Eagle&#39;s Medium</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">10566-016</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;"></td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="width:204px;height:20px;">B) Antibodies and cell lysates</td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">CD31&#160;</td>
			<td style="width:127px;">Abcam</td>
			<td style="width:119px;">ab28364</td>
			<td>1:250 dilution&#160; for Western blotting</td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">CD34</td>
			<td style="width:127px;">Santa Cruz Biotechnology</td>
			<td style="width:119px;">sc-7045</td>
			<td>1:100 dilution for Western blotting</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">&#945;-SMA</td>
			<td style="width:127px;">abcam</td>
			<td style="width:119px;">ab5694</td>
			<td>1:100 dilution for Western blotting</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">&#945;-tubulin</td>
			<td style="width:127px;">abcam</td>
			<td style="width:119px;">ab7291</td>
			<td>1:2500 dilution for Western blotting</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">VEGFR2</td>
			<td style="width:127px;">abcam</td>
			<td style="width:119px;">sc504</td>
			<td>1:100 dilution for Western blotting</td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Human umbilical vein endothelial cell lysate</td>
			<td style="width:127px;">Santa Cruz Biotechnology</td>
			<td style="width:119px;">sc24709&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">Valve interstitial cell lysate</td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td style="width:807px;">Primary cell line cultured from own lab and lysed with RIPA buffer</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;"></td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="width:204px;height:40px;">C) Western blotting and immunostaining</td>
			<td style="width:127px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">10X Tris/Glycine/SDS buffer</td>
			<td style="width:127px;">Biorad</td>
			<td style="width:119px;">161-0772</td>
			<td>Used as running buffer</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">10X Tris/Glycine buffer</td>
			<td style="width:127px;">Biorad</td>
			<td style="width:119px;">161-0771</td>
			<td>Used as transfer buffer</td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Immobilon-FL transfer membrane</td>
			<td style="width:127px;">Merck Millipore</td>
			<td style="width:119px;">IPFL0010</td>
			<td>This is a PVDF transfer membrane that has 45 &#181;m pore size and is mentioned in the protocol as western blot membrane</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">4X Laemmli sample buffer</td>
			<td style="width:127px;">Biorad</td>
			<td style="width:119px;">161-0747</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">2-mercaptoethanol</td>
			<td style="width:127px;">Biorad</td>
			<td style="width:119px;">161-0710</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">10% Criterion TGX precast gel</td>
			<td style="width:127px;">Biorad</td>
			<td style="width:119px;">5671033</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="width:204px;height:42px;">Prolong Gold antifade</td>
			<td style="width:127px;">Thermofisher Scientific</td>
			<td style="width:119px;">P36930</td>
			<td>Used for mounting immunostained coverslips for long term storage</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:204px;height:21px;">Methanol</td>
			<td style="width:127px;">VWR Analytical</td>
			<td style="width:119px;">BDH1135-4LP</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Isolation and Expansion of Endothelial Progenitor Cells: 
A schematic (Figure 1) is provided depicting the overall protocol. The different blood component layers were observed following density gradient centrifugation of human umbilical cord blood with density gradient medium. Upon seeding MNCs onto the collagen-treated plates, the outgrowth of colonies was first observed between Days 5 and 7 (Figure 2<stron...

Watch this Scientific Journal Video about Isolation of Endothelial Progenitor Cells from Human Umbilical Cord Blood at JoVE.com

Isolation of Endothelial Progenitor Cells from Human Umbilical Cord Blood

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 ...

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Research

JoVE Journal

Developmental Biology

11.0K Views.  University of Arkansas. 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.

Video: Isolation of Endothelial Progenitor Cells from Human Umbilical Cord Blood

Direct Induction of Human Neural Stem Cells from Peripheral Blood Hematopoietic Progenitor Cells

Human disease specific neuronal cultures are essential for generating in vitro models for human neurological diseases. However, the lack of access to primary human adult neural cultures raises unique challenges. Recent developments in induced pluripotent stem cells (iPSC) provides an alternative approach to derive neural cultures from skin fibroblasts through patient specific iPSC, but this process is labor intensive, requires special expertise and large amounts of resources, and can take several months. This prevents the wide application of this technology to the study of neurological diseases. To overcome some of these issues, we have developed a method to derive neural stem cells directly from human adult peripheral blood, bypassing the iPSC derivation process. Hematopoietic progenitor cells enriched from human adult peripheral blood were cultured in vitro and transfected with Sendai virus vectors containing transcriptional factors Sox2, Oct3/4, Klf4, and c-Myc. The transfection results in morphological changes in the cells which are further selected by using human neural progenitor medium containing basic fibroblast growth factor (bFGF) and vascular endothelial growth factor (VEGF). The resulting cells are characterized by the expression for neural stem cell markers, such as nestin and SOX2. These neural stem cells could be further differentiated to neurons, astroglia and oligodendrocytes in specified differentiation media. Using easily accessible human peripheral blood samples, this method could be used to derive neural stem cells for further differentiation to neural cells for in vitro modeling of neurological disorders and may advance studies related to the pathogenesis and treatment of those diseases.

A method was developed to directly derive human neural stem cells from hematopoietic progenitor cells enriched from peripheral blood cells.

Human disease specific neuronal cultures are essential for generating in vitro models for human neurological diseases. However, the lack of access to ...

Isolation of Neural Stem/Progenitor Cells from the Periventricular Region of the Adult Rat and Human Spinal Cord

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of multipotent, self-renewing, and expandable neural stem cells. In serum conditions, these multipotent NSPCs will differentiate, generating neurons, astrocytes, and oligodendrocytes. The harvested tissue is enzymatically dissociated in a papain-EDTA solution and then mechanically dissociated and separated through a discontinuous density gradient to yield a single cell suspension which is plated in neurobasal medium supplemented with epidermal growth factor (EGF), basic fibroblast growth factor (bFGF), and heparin. Adult rat spinal cord NSPCs are cultured as free-floating neurospheres and adult human spinal cord NSPCs are grown as adherent cultures. Under these conditions, adult spinal cord NSPCs proliferate, express markers of precursor cells, and can be continuously expanded upon passage. These cells can be studied in vitro in response to various stimuli, and exogenous factors may be used to promote lineage restriction to examine neural stem cell differentiation. Multipotent NSPCs or their progeny can also be transplanted into various animal models to assess regenerative repair.

The adult mammalian spinal cord contains neural stem/progenitor cells (NSPCs) that can be isolated and expanded in culture. This protocol describes the harvesting, isolation, culture, and passaging of NSPCs generated from the periventricular region of the adult spinal cord from the rat and from human organ transplant donors.

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of ...

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, ...

Isolating Mesangiogenic Progenitor Cells (MPCs) from Human Bone Marrow

In a research study aimed to isolate human bone marrow (hBM)-derived Mesenchymal Stromal Cells (MSCs) for clinical applications, we identified a novel cell population specifically selected for growth in human serum supplemented medium. These cells are characterized by morphological, phenotypic, and molecular features distinct from MSCs and we named them Mesodermal Progenitor Cells (MPCs). MPCs are round, with a thick highly refringent core region; they show strong, trypsin resistant adherence to plastic. Failure to expand MPCs directly revealed that they are slow in cycling. This is as also suggested by Ki-67 negativity. On the other hand, culturing MPCs in standard medium designed for MSC expansion, gave rise to a population of exponentially growing MSC-like cells. Besides showing mesenchymal differentiation capacity MPCs retained angiogenic potential, confirming their multiple lineage progenitor nature. Here we describe an optimized highly reproducible protocol to isolate and characterize hBM-MPCs by flow cytometry (CD73, CD90, CD31, and CD45), nestin expression, and F-actin organization. Protocols for mesengenic and angiogenic differentiation of MPCs are also provided. Here we also suggest a more appropriate nomenclature for these cells, which has been re-named as "Mesangiogenic Progenitor Cells".

Isolating Mesangiogenic Progenitor Cells MPCs from Human Bone Marrow

Here we describe an optimized, highly reproducible protocol to isolate Mesodermal Progenitor Cells (MPCs) from human bone marrow (hBM). MPCs were characterized by flow cytometry and nestin expression. They showed the ability to give rise to exponentially growing MSC-like cell cultures while retaining their angiogenic potential.

In a research study aimed to isolate human bone marrow (hBM)-derived Mesenchymal Stromal Cells (MSCs) for clinical applications, we identified a novel ...

Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly gained attention because they do not pose any ethical or moral concerns. Current methods to isolate MSCs from UC yield low amounts of cells with variable proliferation potentials. Since UC is an anatomically-complex organ, differences in MSC properties may be due to the differences in the anatomical regions of their isolation. In this study, we first dissected the cord/placenta samples into three discrete anatomical regions: UC, cord-placenta junction (CPJ), and fetal placenta (FP). Second, two distinct zones, cord lining (CL) and Wharton&#39;s jelly (WJ), were separated. The explant culture technique was then used to isolate cells from the four sources. The time required for the primary culture of cells from the explants varied depending on the source of the tissue. Outgrowth of the cells occurred within 3 - 4 days of the CPJ explants, whereas growth was observed after 7 - 10 days and 11 - 14 days from CL/WJ and FP explants, respectively. The isolated cells were adherent to plastic and displayed fibroblastoid morphology and surface markers, such as CD29, CD44, CD73, CD90, and CD105, similarly to bone marrow (BM)-derived MSCs. However, the colony-forming efficiency of the cells varied, with CPJ-MSCs and WJ-MSCs showing higher efficiency than BM-MSCs. MSCs from all four sources differentiated into adipogenic, chondrogenic, and osteogenic lineages, indicating that they were multipotent. CPJ-MSCs differentiated more efficiently in comparison to other MSC sources. These results suggest that the CPJ is the most potent anatomical region and yields a higher number of cells, with greater proliferation and self-renewal capacities in vitro. In conclusion, the comparative analysis of the MSCs from the four sources indicated that CPJ is a more promising source of MSCs for cell therapy, regenerative medicine, and tissue engineering.

Here, we present a protocol for the dissection of human umbilical cord (UC) and fetal placenta sample into cord lining (CL), Wharton&#39;s jelly (WJ), cord-placenta junction (CPJ), and fetal placenta (FP) for the isolation and characterization of mesenchymal stromal cells (MSCs) using the explant culture technique.

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly ...

Chondrogenic Pellet Formation from Cord Blood-derived Induced Pluripotent Stem Cells

Human articular cartilage lacks the ability to repair itself. Cartilage degeneration is thus treated not by curative but by conservative treatments. Currently, efforts are being made to regenerate damaged cartilage with ex vivo expanded chondrocytes or bone marrow-derived mesenchymal stem cells (BMSCs). However, the restricted viability and instability of these cells limit their application in cartilage reconstruction. Human induced pluripotent stem cells (hiPSCs) have received scientific attention as a new alternative for regenerative applications. With unlimited self-renewal ability and multipotency, hiPSCs have been highlighted as a new replacement cell source for cartilage repair. However, obtaining a high quantity of high-quality chondrogenic pellets is a major challenge to their clinical application. In this study, we used embryoid body (EB)-derived outgrowth cells for chondrogenic differentiation. Successful chondrogenesis was confirmed by PCR and staining with alcian blue, toluidine blue, and antibodies against collagen types I and II (COL1A1 and COL2A1, respectively). We provide a detailed method for the differentiation of cord blood mononuclear cell-derived iPSCs (CBMC-hiPSCs) into chondrogenic pellets.

Here, we propose a protocol for chondrogenic differentiation from cord blood mononuclear cell-derived human induced pluripotent stem cells.

Human articular cartilage lacks the ability to repair itself. Cartilage degeneration is thus treated not by curative but by conservative treatments. ...

Induction of Endothelial Differentiation in Cardiac Progenitor Cells Under Low Serum Conditions

Cardiac progenitor cells (CPCs) may have therapeutic potential for cardiac regeneration after injury. In the adult mammalian heart, intrinsic CPCs are extremely scarce, but expanded CPCs could be useful for cell therapy. A prerequisite for their use is their ability to differentiate in a controlled manner into the various cardiac lineages using defined and efficient protocols. In addition, upon in vitro expansion, CPCs isolated from patients or preclinical disease models may offer fruitful research tools for the investigation of disease mechanisms.
Current studies use different markers to identify CPCs. However, not all of them are expressed in humans, which limits the translational impact of some preclinical studies. Differentiation protocols that are applicable irrespective of the isolation technique and marker expression will allow for the standardized expansion and priming of CPCs for cell therapy purpose. Here we describe that the priming of CPCs under a low fetal bovine serum (FBS) concentration and low cell density conditions facilitates the endothelial differentiation of CPCs. Using two different subpopulations of mouse and rat CPCs, we show that laminin is a more suitable substrate than fibronectin for this purpose under the following protocol: after culturing for 2 - 3 days in medium including supplements that maintain multipotency and with 3.5% FBS, CPCs are seeded on laminin at &#60;60% confluence and cultured in supplement-free medium with low concentrations of FBS (0.1%) for 20 - 24 hours before differentiation in endothelial differentiation medium. Because CPCs are a heterogeneous population, serum concentrations and incubation times may need to be adjusted depending on the properties of the respective CPC subpopulation. Considering this, the technique can be applied to other types of CPCs as well and provides a useful method to investigate the potential and mechanisms of differentiation and how they are affected by disease when using CPCs isolated from respective disease models.

This protocol describes an endothelial differentiation technique for cardiac progenitor cells. It particularly focuses on how serum concentration and cell-seeding density affect the endothelial differentiation potential.

Cardiac progenitor cells (CPCs) may have therapeutic potential for cardiac regeneration after injury. In the adult mammalian heart, intrinsic CPCs are ...

Isolation and Characterization of Human Umbilical Cord-derived Mesenchymal Stem Cells from Preterm and Term Infants

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally isolated and characterized from bone marrow (BM), then acquired from tissues including adipose tissue, synovium, skin, dental pulp, and fetal appendages such as placenta, umbilical cord blood (UCB), and umbilical cord (UC). MSCs are a heterogeneous cell population with the capacity for (1) adherence to plastic in standard culture conditions, (2) surface marker expression of CD73+/CD90+/CD105+/CD45-/CD34-/CD14-/CD19-/HLA-DR- phenotypes, and (3) trilineage differentiation into adipocytes, osteocytes, and chondrocytes, as currently defined by the International Society for Cellular Therapy (ISCT). Although BM is the most widely used source of MSCs, the invasive nature of BM aspiration ethically limits its accessibility. Proliferation and differentiation capacity of MSCs obtained from BM generally decline with the age of the donor. In contrast, fetal MSCs obtained from UC have advantages such as vigorous proliferation and differentiation capacity. There is no ethical concern for UC sampling, as it is typically regarded as medical waste. Human UC starts to develop with continuing growth of the amniotic cavity at 4-8 weeks of gestation and keeps growing until reaching 50-60 cm in length, and it can be isolated during the whole newborn delivery period. To gain insight into the pathophysiology of intractable diseases, we have used UC-derived MSCs (UC-MSCs) from infants delivered at various gestational ages. In this protocol, we describe the isolation and characterization of UC-MSCs from fetuses/infants at 19-40 weeks of gestation.

Human umbilical cord (UC) can be obtained during the perinatal period as a result of preterm, term, and postterm delivery. In this protocol, we describe the isolation and characterization of UC-derived mesenchymal stem cells (UC-MSCs) from fetuses/infants at 19-40 weeks of gestation.

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally ...

Generation of 3D Skin Organoid from Cord Blood-derived Induced Pluripotent Stem Cells

The skin is the body&#8217;s largest organ and has many functions. The skin acts as a physical barrier and protector of the body and regulates bodily functions. Biomimetics is the imitation of the models, systems, and elements of nature for the purpose of solving complex human problems1. Skin biomimetics is a useful tool for in vitro disease research and in vivo regenerative medicine. Human induced pluripotent stem cells (iPSCs) have the characteristic of unlimited proliferation and the ability of differentiation to three germ layers. Human iPSCs are generated from various primary cells, such as blood cells, keratinocytes, fibroblasts, and more. Among them, cord blood mononuclear cells (CBMCs) have emerged as an alternative cell source from the perspective of allogeneic regenerative medicine. CBMCs are useful in regenerative medicine because human leukocyte antigen (HLA) typing is essential to the cell banking system. We provide a method for the differentiation of CBMC-iPSCs into keratinocytes and fibroblasts and for generation of a 3D skin organoid. CBMC-iPSC-derived keratinocytes and fibroblasts have characteristics similar to a primary cell line. The 3D skin organoids are generated by overlaying an epidermal layer onto a dermal layer. By transplanting this 3D skin organoid, a humanized mice model is generated. This study shows that a 3D human iPSC-derived skin organoid may be a novel, alternative tool for dermatologic research in vitro and in vivo.

We propose a protocol that shows how to differentiate induced pluripotent stem cell-derived keratinocytes and fibroblasts and generate a 3D skin organoid, using these keratinocytes and fibroblasts. This protocol contains an additional step of generating a humanized mice model. The technique presented here will improve dermatologic research.

The skin is the body&#8217;s largest organ and has many functions. The skin acts as a physical barrier and protector of the body and regulates bodily ...

In Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...