This Project was supported by P01HL095489. A.Y.H was supported by T32HL066987.

NOTE: Human neutrophils were isolated from venous blood from discarded white blood cell filters obtained from the Blood Bank Lab at the Boston Children&#39;s Hospital. Blood donors were unidentifiable, and there was no interaction with living individuals or knowledge of identifiable personal information. Therefore, this work is not classified as human subjects research under the HHS human subjects regulations (45 CFR Part 46). The Boston Children&#39;s Hospital Institutional Review Board (IRB) approved the protocol.
1. Layering the gradient
<ol>
	<li>After sterilizing the buffy coat or whole blood packaging and the laminar hood, divide the blood into 50 mL tubes with 10 mL of blood in each tube.</li>
	<li>Bring the volume up to 35 mL with 5% fetal bovine serum (FBS)/Hank&#39;s Balanced Salt Solution (HBSS) to dilute the blood for a cleaner gradient. 
	NOTE: When using a leukapheresis membrane, cells can be flushed out using a 60 mL syringe and 30 mL of 5% FBS/HBSS per filter. Dilution is unnecessary when working with freshly drawn whole blood.</li>
	<li>Close the 50 mL tube lid, mix it several times by inversion, and keep it upside down to have the bottom devoid of RBCs.</li>
	<li>Add 10 mL of density gradient medium (see the Table of Materials) directly beneath the blood. Ensure that the medium and the blood do not mix and that the interface is sharp (Figure 1A). 
	NOTE: This step is crucial. Ensure that the density gradient solution is at room temperature (RT) and well-mixed before each gradient. The first milliliter of the density gradient medium needs to be carefully and steadily layered as slowly as possible. Having the electronic pipette speed set to low is recommended.&#160;</li>
	<li>Gently place the tube in a centrifuge without disturbing the gradient and spin at 400 &#215; g for 30 min at RT, making sure to disable the brake. Observe how the spun gradient separates into a top serum/plasma layer, a middle white ring of peripheral blood mononuclear cells (PBMCs), a cloudy density gradient medium layer, and a bottom pellet consisting of a white, thin neutrophil band on top of the RBCs (Figure 1B). 
	NOTE: A cloudy or opaque side of the tube after centrifugation can suggest that the cells (neutrophils) are activated and may not be suitable to use.</li>
	<li>Remove the PBMCs first by emerging the suction pipette directly into the PBMC layer. Be sure to aspirate it completely while the serum/plasma layer decreases as the ring is removed. Scrape the side of the tube where cells pelleted with the suction pipette to maximize the removal of the PMBC. Carefully remove the cloudy density gradient medium layer between the PBMC ring and the neutrophil/RBC pellet. 
	&#8203;NOTE: Scraping the pelleted cells on the side of the tube increases the purity of the isolation considerably. Be careful not to aspirate the pellet, as most neutrophils are sitting directly on top of the RBCs.</li>
</ol>
2. Sedimentation of erythrocytes (RBCs)
<ol>
	<li>Using a 10 mL pipette, transfer the neutrophil/RBC pellet into a clean tube. Do not pipette up and down. Add 5% FBS/HBSS to a final volume of 25 mL. Gently mix by inversion. 
	NOTE: Performing the RBC sedimentation after the PBMC removal improves yield and decreases activation10.</li>
	<li>Directly add 25 mL of prewarmed (37 &#176;C) 3% Dextran/0.9% NaCl/H2O into the tube containing the diluted neutrophil/RBC pellet and mix gently by inversion. Place the tube on a level, non-vibrating surface for 15 min (Figure 1C). 
	NOTE: Longer sedimentation reduces RBC contamination but also decreases the yield. Furthermore, prolonged dextran exposure may lead to neutrophil activation or cell death11.</li>
	<li>Gently bring the tube back into the hood (for sterile isolation). By only slightly immersing the pipette in the liquid, collect the top layer (~30 mL) following the liquid surface downwards. 
	NOTE: If the sedimentation produces a sharp interface between the medium and the RBCs, the RBC pellet is smaller, or if a higher yield is desired, collect up to 35 mL.</li>
	<li>Spin the tube (400 &#215; g, 10 min, RT, using low brake (3)), resulting in a red pellet with no particles floating in the media.</li>
</ol>
3. Lysis of the residual RBCs
<ol>
	<li>Gently aspirate the supernatant without disrupting the pellet.</li>
	<li>Add 25 mL of sterile ultrapure water directly into the tube and gently mix by inverting for 28 s to lyse the RBCs. Do not use a pipet to resuspend the pellet. 
	NOTE: Do not exceed 30 s as prolonged hypotonic conditions can activate and lead to neutrophil death12.</li>
	<li>Immediately add 25 mL of sterile 1.8% NaCl/H2O into the tube and gently mix by inverting to bring the solution back to isotonic conditions. 
	NOTE: The solution should be red but without turbidity.</li>
	<li>Spin down at 200 &#215; g for 3-5 min with low brake (level 3) to minimize RBCs and platelet sedimenting together with the neutrophils (Figure 1D and Figure 2)13,14. 
	NOTE: The pellet should be white with a minimal RBC layer on top, which can be gently removed while the supernatant is aspirated.</li>
	<li>Resuspend neutrophils by pipetting the culture medium (10% FBS/RPMI1640) directly on the pellet, but do not pipette up and down. Rock the tube horizontally from side to side to minimize cell activation. 
	NOTE: Cells should be kept at a concentration of ~2 &#215; 106 cells/mL, as higher density will lead to increased cell activation/death15. For the same reason, the cell pellet should be resuspended as soon as possible.</li>
	<li>If cell aggregation or clumping is observed, filter the cell suspension using a 70 &#181;m mesh to discard clumped neutrophils.</li>
</ol>
4. Determining neutrophil isolation quality
<ol>
	<li>Stain the cells with markers specific for neutrophils (CD66b, CD11b), eosinophils (CD193) (Figure 3), and an activation marker such as CD62L (Figure 4). Acquire 20,000 cells by flow cytometry. Analyze cell purity and activation using the gating strategies proposed in Figure 3 and Figure 4. 
	NOTE: The quality of the neutrophil preparation can be assessed using a 3% acetic acid-methylene blue solution to visualize the unique lobed nucleus of neutrophils.</li>
	<li>Determine cell viability using Annexin V/propidium iodide (PI) (Figure 5). 
	NOTE: Trypan blue staining can be used to evaluate cell viability.</li>
</ol>

<ol>
	<li>Rosales, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil:+A+cell+with+many+roles+in+inflammation+or+several+cell+types.">Neutrophil: A cell with many roles in inflammation or several cell types.</a> Frontiers in Physiology. 9, 113 (2018).</li><li>Amulic, B., Cazalet, C., Hayes, G. L., Metzler, K. D., Zychlinsky, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil+function:+from+mechanisms+to+disease.">Neutrophil function: from mechanisms to disease.</a> Annual Review of Immunology. 30, 459-489 (2012).</li><li>Stoller, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murray+&amp;+Nadel's+textbook+of+respiratory+medicine,+6th+edition.">Murray &amp; Nadel's textbook of respiratory medicine, 6th edition.</a> Annals of the American Thoracic Society. 12 (8), 1257-1258 (2015).</li><li>Dancey, J. T., Deubelbeiss, K. A., Harker, L. A., Finch, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil+kinetics+in+man.">Neutrophil kinetics in man.</a> Journal of Clinical Investigation. 58 (3), 705-715 (1976).</li><li>Manz, M. G., Boettcher, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emergency+granulopoiesis.">Emergency granulopoiesis.</a> Nature Reviews. Immunology. 14 (5), 302-314 (2014).</li><li>Summers, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil+kinetics+in+health+and+disease.">Neutrophil kinetics in health and disease.</a> Trends in Immunology. 31 (8), 318-324 (2010).</li><li>Pedruzzi, E., Fay, M., Elbim, C., Gaudry, M., Gougerot-Pocidalo, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differentiation+of+PLB-985+myeloid+cells+into+mature+neutrophils,+shown+by+degranulation+of+terminally+differentiated+compartments+in+response+to+N-formyl+peptide+and+priming+of+superoxide+anion+production+by+granulocyte-macrophage+colony-stimulating+factor.">Differentiation of PLB-985 myeloid cells into mature neutrophils, shown by degranulation of terminally differentiated compartments in response to N-formyl peptide and priming of superoxide anion production by granulocyte-macrophage colony-stimulating factor.</a> British Journal of Haematology. 117 (3), 719-726 (2002).</li><li>Tucker, K. A., Lilly, M. B., Heck, L., Rado, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+a+new+human-diploid+myeloid-leukemia+cell-line+(Plb-985)+with+granulocytic+and+monocytic+differentiating+capacity.">Characterization of a new human-diploid myeloid-leukemia cell-line (Plb-985) with granulocytic and monocytic differentiating capacity.</a> Blood. 70 (2), 372-378 (1987).</li><li>Hsu, A. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inducible+overexpression+of+zebrafish+microRNA-722+suppresses+chemotaxis+of+human+neutrophil+like+cells.">Inducible overexpression of zebrafish microRNA-722 suppresses chemotaxis of human neutrophil like cells.</a> Molecular Immunology. 112, 206-214 (2019).</li><li>Kremserova, S., Nauseef, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+human+neutrophils+from+venous+blood.">Isolation of human neutrophils from venous blood.</a> Methods in Molecular Biology. 2087, 33-42 (2020).</li><li>Quach, A., Ferrante, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+application+of+dextran+sedimentation+as+an+initial+step+in+neutrophil+purification+promotes+their+stimulation,+due+to+the+presence+of+monocytes.">The application of dextran sedimentation as an initial step in neutrophil purification promotes their stimulation, due to the presence of monocytes.</a> Journal of Immunological Research. 2017, 1254792 (2017).</li><li>Thorson, L. M., Turkalj, A., Hung, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+evaluation+of+neutrophil+viability+after+exposure+to+a+hypotonic+medium.">In vitro evaluation of neutrophil viability after exposure to a hypotonic medium.</a> Nuclear Medicine Communications. 16 (7), 615-620 (1995).</li><li>Dhurat, R., Sukesh, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+and+methods+of+preparation+of+platelet-rich+plasma:+a+review+and+author's+perspective.">Principles and methods of preparation of platelet-rich plasma: a review and author's perspective.</a> Journal of Cutaneous and Aesthetetic Surgery. 7 (4), 189-197 (2014).</li><li>Etulain, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimised+protocol+for+platelet-rich+plasma+preparation+to+improve+its+angiogenic+and+regenerative+properties.">An optimised protocol for platelet-rich plasma preparation to improve its angiogenic and regenerative properties.</a> Scientific Reports. 8 (1), 1513 (2018).</li><li>Hannah, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Constitutive+neutrophil+apoptosis+in+culture+is+modulated+by+cell+density+independently+of+beta2+integrin-mediated+adhesion.">Constitutive neutrophil apoptosis in culture is modulated by cell density independently of beta2 integrin-mediated adhesion.</a> FEBS Letters. 421 (2), 141-146 (1998).</li><li>Hsu, A. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotypical+microRNA+screen+reveals+a+noncanonical+role+of+CDK2+in+regulating+neutrophil+migration.">Phenotypical microRNA screen reveals a noncanonical role of CDK2 in regulating neutrophil migration.</a> Proceedings of the National Acadermy of Sciences of the United States of America. 116 (37), 18561-18570 (2019).</li><li>Martin, S. J., Bradley, J. G., Cotter, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=HL-60+cells+induced+to+differentiate+towards+neutrophils+subsequently+die+via+apoptosis.">HL-60 cells induced to differentiate towards neutrophils subsequently die via apoptosis.</a> Clinical and Experimental Immunology. 79 (3), 448-453 (1990).</li><li>Hauert, A. B., Martinelli, S., Marone, C., Niggli, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differentiated+HL-60+cells+are+a+valid+model+system+for+the+analysis+of+human+neutrophil+migration+and+chemotaxis.">Differentiated HL-60 cells are a valid model system for the analysis of human neutrophil migration and chemotaxis.</a> International Journal of Biochemistry and Cell Biology. 34 (7), 838-854 (2002).</li><li>Blanter, M., Gouwy, M., Struyf, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studying+neutrophil+function+in+vitro:+Cell+models+and+environmental+factors.">Studying neutrophil function in vitro: Cell models and environmental factors.</a> Journal of Inflammation Research. 14, 141-162 (2021).</li><li>Kambara, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gasdermin+D+exerts+anti-inflammatory+effects+by+promoting+neutrophil+death.">Gasdermin D exerts anti-inflammatory effects by promoting neutrophil death.</a> Cell Reports. 22 (11), 2924-2936 (2018).</li><li>Loison, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Proteinase+3-dependent+caspase-3+cleavage+modulates+neutrophil+death+and+inflammation.">Proteinase 3-dependent caspase-3 cleavage modulates neutrophil death and inflammation.</a> Journal of Clinical Investigation. 124 (10), 4445-4458 (2014).</li><li>Siemsen, D. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil+isolation+from+nonhuman+species.">Neutrophil isolation from nonhuman species.</a> Methods in Molecular Biology. 1124, 19-37 (2014).</li><li>Hasenberg, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+immunomagnetic+negative+enrichment+of+neutrophil+granulocytes+from+murine+bone+marrow+for+functional+studies+in+vitro+and+in+vivo.">Rapid immunomagnetic negative enrichment of neutrophil granulocytes from murine bone marrow for functional studies in vitro and in vivo.</a> PLoS One. 6 (2), 17314 (2011).</li><li>Calzetti, F., Tamassia, N., Arruda-Silva, F., Gasperini, S., Cassatella, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+importance+of+being+"pure"+neutrophils.">The importance of being "pure" neutrophils.</a> Journal of Allergy and Clinical Immunology. 139 (1), 352-355 (2017).</li></ol>

The authors declare that the research was conducted in the absence of any conflict of interest.

Due to the short life span, terminal differentiation status, and lytic content of neutrophils, studying these cells has always been a challenge. Apart from utilizing mouse models or cells from patient cohorts, cell lines are useful tools to help study neutrophil biology16. However, neutrophil-like cell lines cannot completely reiterate all aspects of neutrophil biology, adding an extra layer of difficulty in studying these cells. The most commonly used in vitro model is the HL-60 cell line, which can be differentiated into neutrophil-like cells by treatment with dimethylsulfoxide or retinoic acid17,18. Although these cells are useful in the study of migration and respiratory burst, they are not suitable for studying the microbiocidal activity of neutrophils. Other cell lines exist (PLB-98, NB4), and they are also associated with their set of limitations19. 
It is pivotal to validate with primary human neutrophils observations made with mouse disease models and cell lines. Neutrophils cannot be cryopreserved efficiently and thus are often freshly isolated from whole blood or buffy coats obtained from donors and immediately processed. Once isolated, the cells start to undergo a complex form of spontaneous death, regulated by oxidation, cytoplasmic caspases, and proteases found in neutrophil granules20,21. Improper isolation methods or techniques can lead to the activation of neutrophils, only accelerating cell demise. It is imperative to have a reliable and consistent method to obtain pure and high-quality neutrophils from donors. 
There have been many methods published on human neutrophil isolation10,22. They mainly fall into two categories, with some shared strategies. The first category is antibody-based, being either through positive or negative selection. Positive selection would label neutrophils directly, therefore providing a highly pure cell population, although it also leads to rapid cell activation, cell death, as well as unwanted tagging of neutrophils23. Negative selection, though leaving the cells unlabeled and giving a very pure population, accelerated neutrophil death, although the precise mechanism is unknown (Figure 5). Whether gene or protein expression is also altered after positive or negative selection needs to be further investigated. Moreover, due to the amount of antibody needed to deplete the other types of cells, these methods cannot output large amounts of neutrophils. However, antibody-based assays can still be used for short-term culture and experiments on a smaller scale and are methods of choice for experiments requiring very high cell purity, such as gene and protein expression studies. 
The second type of isolation method is gradient- and density-based. It usually involves Percoll, Ficoll-Paque, or other polysaccharide/polyvinylpyrrolidone components and utilizes centrifugation force to separate different types of blood cells based on cell density. These methods often are complemented with the sedimentation of red blood cells by dextran. These methods can handle larger scales of starting material and can achieve high purity as well. One caveat of density-based isolation is the inefficient separation of other far less abundant granulocytes (mostly eosinophils) from neutrophils, and it is, therefore, the major limitation of the protocol presented, as even the presence of small cell contamination can affect neutrophil response24.
Presented here is a summarized method based on gradient isolation, refining previous methods10,22. We utilize the current understating of neutrophil specificities to reliably isolate pure human neutrophils with limited residual platelet and RBCs, preventing neutrophil activation and accelerated death. The most crucial step is the layering of the gradient, which is much more effectively obtained by adding the density gradient medium underneath the blood to obtain a sharp interface. A quick examination of the PBMC ring and the gradient after centrifugation can reveal possible contamination, activation, and low yields. When working with a leukapheresis membrane or buffy coat, diluting the blood is important as excessive cell density would lead to cell aggregation, leading to impurities and cell activation. 
This protocol should be completed within 2 h to ensure cell freshness, and steps involving the density gradient medium, dextran, and lysis be done immediately, as exposure to these solutions can alter neutrophils. With this protocol, the expected yield of neutrophils is at least ~10 million/10 mL of whole blood and at least ~60 million/10 mL of buffy coat. Evaluation of isolation quality should be done as follows: activated neutrophils should be less than 10%, lymphocyte contamination lower than 5%, minimal eosinophil (same side scatter but lower forward scatter population), and cell viability should be over 90%. Lower purity can result from improper layering or storage of the density gradient medium or due to the quality and freshness of the starting blood product.

The innate immune system is comprised of many cell types that maintain immune homeostasis and pathogen clearance along with many other physiological functions. Neutrophils comprise the largest pool of white blood cells in human circulation1. Most mature neutrophils are stored in the bone marrow, which is the site of generation for new neutrophils, also called granulopoiesis. In the bone marrow, granulocyte progenitors exit the cell cycle and terminally differentiate, acquiring their characteristic segmented nuclei and granules2. Under inflammatory conditions, in response to chemokines, cytokines, and damage-associated and pathogen-associated molecular patterns, neutrophils are mobilized from the bloodstream and out of the bone marrow to perform a wide array of functions. These include cytokine secretion, direct phagocytosis of the pathogen, the release of reactive oxygen species, degranulation of antimicrobial proteins, and formation of neutrophil extracellular traps.
The molecules used by neutrophils to fight infection are toxic to the microbes and the host. Thus, the life span and the proper removal of aging/dying neutrophils are highly regulated, and they have a limited life span in circulation (&#60;48 h)3. Due to this short survival, the human body produces on average 100 billion new neutrophils&#160;every day to maintain population homeostasis4. Emergency granulopoiesis can further increase the release of neutrophils, both mature and immature, in the blood during inflammation and infection5. The importance of neutrophils in the innate immune response is highlighted by patients with acquired or congenital neutropenia, who are susceptible to bacterial and fungal infections6.
Many challenges arise when studying neutrophil biology and their roles in the immune response due to their nature, including their short survival and cytotoxic content. Neutrophil-like cell lines have been commonly differentiated from human promyelocytic leukemia HL-60 cells and PLB-985 cells7,8. Although they can display neutrophil-like morphology and perform chemotaxis, these cell lines cannot fully recapitulate the biology of neutrophils. In vitro assays using these cell lines are also unable to recapitulate in vivo experiments. Furthermore, the differentiation of these cells needs to be induced and could be adversely affected by gene manipulation before differentiation.
Recently, methods have been developed to circumvent these issues by using inducible promoters to modulate gene expression post differentiation in HL-60 cells9. Even with such tools, primary human PMNs are required to validate targets using pharmacological approaches. Thus, it is imperative to obtain pure and inactivated neutrophils isolated from blood to validate the findings of cell line and animal models. This paper presents a revised PMN isolation protocol in which the pros and cons of current methods were evaluated10. A combination was devised consisting of gradient centrifugation to separate PMNs from other immune cells, short dextran-based sedimentation to remove the bulk of RBCs, rapid residual RBC lysis via osmotic pressure, and low-speed centrifugation to remove platelet contamination.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3% Acetic Acid with Methylene Blue</td>
			<td>Stemcell technologies</td>
			<td>#07060</td>
			<td></td>
		</tr>
		<tr>
			<td>Attune NxT</td>
			<td>invitrogen</td>
			<td>A24858</td>
			<td>FACS analyzer</td>
		</tr>
		<tr>
			<td>Cd11b-PE</td>
			<td>biolegend</td>
			<td>301305</td>
			<td></td>
		</tr>
		<tr>
			<td>CD14-BV421</td>
			<td>biolegend</td>
			<td>367144</td>
			<td></td>
		</tr>
		<tr>
			<td>CD193-BV605</td>
			<td>biolegend</td>
			<td>310716</td>
			<td></td>
		</tr>
		<tr>
			<td>CD19-PerCP</td>
			<td>biolegend</td>
			<td>302228</td>
			<td></td>
		</tr>
		<tr>
			<td>CD3-PerCP</td>
			<td>biolegend</td>
			<td>300326</td>
			<td></td>
		</tr>
		<tr>
			<td>CD41-APC</td>
			<td>biolegend</td>
			<td>303710</td>
			<td></td>
		</tr>
		<tr>
			<td>Cd45-APC</td>
			<td>biolegend</td>
			<td>103111</td>
			<td></td>
		</tr>
		<tr>
			<td>CD62L-PE</td>
			<td>biolegend</td>
			<td>304802</td>
			<td></td>
		</tr>
		<tr>
			<td>CD66b-FITC</td>
			<td>biolegend</td>
			<td>305103</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5810R</td>
			<td>Eppendorf</td>
			<td>22625101</td>
			<td>Centrifuge</td>
		</tr>
		<tr>
			<td>Dextran</td>
			<td>Fisher</td>
			<td>BP1580</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>R&#38;D systems</td>
			<td>S11150H</td>
			<td>complement inactivation of FBS is recommended</td>
		</tr>
		<tr>
			<td>FITC Annexin V Apoptosis Detection Kit I</td>
			<td>BD</td>
			<td>556547</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks balanced salt solution (-CaCl2), (-MgCl2) (-MgSO4)</td>
			<td>Gibco</td>
			<td>14175-095</td>
			<td>HBSS without Ca2+/Mg2+ is advised&#160; as they have been shown to lead to neutrohpil activation</td>
		</tr>
		<tr>
			<td>Lymphoprep</td>
			<td>Stemcell technologies</td>
			<td>#07801</td>
			<td>density gradient medium</td>
		</tr>
		<tr>
			<td>MACSxpress Whole Blood Neutrophil Isolation Kit, human</td>
			<td>Miltenyi</td>
			<td>130-104-434</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI-1640</td>
			<td>Gibco</td>
			<td>11875093</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>sigma</td>
			<td>71376</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue Solution, 0.4%</td>
			<td>Thermofisher</td>
			<td>15250061</td>
			<td></td>
		</tr>
		<tr>
			<td>ultrapure water</td>
			<td>KD medical</td>
			<td>RGF-3410</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation of human neutrophils from whole blood and buffy coats

Neutrophils (PMNs) are the most abundant leukocytes in human circulation, ranging from 40 to 70% of total blood leukocytes. They are the first cells recruited at the site of inflammation via rapid extravasation through vessels. There, neutrophils perform an array of functions to kill invading pathogens and mediate immune signaling. Freshly purified neutrophils from human blood are the model of choice for study, as no cell line fully replicates PMN functions and biology. However, neutrophils are short-lived, terminally differentiated cells and are highly susceptible to activation in response to physical (temperature, centrifugation speed) and biological (endotoxin, chemo- and cytokines) stimuli. Therefore, it is crucial to follow a standardized, reliable, and fast method to obtain pure and non-activated cells. This protocol presents an updated protocol combining density gradient centrifugation, red blood cell (RBC) sedimentation, and RBC lysis to obtain high PMN purity and minimize cell activation. Furthermore, methods to assess neutrophil isolation quality, viability, and purity are also discussed.

When using density gradient to purify neutrophils, it is critical for the interface between the blood and the density gradient medium to be as sharp as possible, and that a distinct layer separation remains after centrifugation (step 1.4). Following RBC lysis, the buffer should be a clear red and not turbid (step 3.3). If the preparation is cloudy, a second round of lysis (step 3) may be required, although this may affect neutrophil survival (Figure 1). Following lysis, low-speed centrifugation (200 &#215; g) is recommended when purity is prioritized, as it reduces platelet contamination considerably. However, high-speed centrifugation (400 &#215; g) increases the yield at the expense of purity (step 3.4, Figure 2). After neutrophil isolation, fluorescence-activated cell sorting can be used to assess isolation purity (step 4.1) and should be chosen over microscopy. Although the FSC/SSC distribution of cells alone provides an estimate of the cell isolation quality (Figure 3A), the use of specific cell markers should be preferred. In this case, the most common contaminating cell populations are stained with specific antibodies together with CD66b, specifically expressed on granulocytes. CD45 staining is used to distinguish leukocytes (CD45+) and red blood cells and platelets (CD45-).
Other contaminants include lymphocytes (CD3+ or CD19+, Figure 3F), monocytes (CD14+, Figure 3D), and eosinophils (CD193+, Figure 3G). CD11b is an integrin expressed on the myeloid lineage; neutrophils and monocytes are CD11b+, whereas lymphocytes are CD11b- (Figure 3C). As neutrophil activation can affect downstream experiments, the expression of CD62L should be assessed; neutrophils become CD62L- once activated (step 4.1). The peptide fMLP can be used as a positive control for CD62L shedding (Figure 4). It is also important to evaluate the health of neutrophils before performing assays; neutrophils have a relatively short half-life, and activation can further shorten it (step 4.2). A standard Annexin V and PI staining can give information on the live/dead status of the neutrophil culture at designated times (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62837/62837fig01.jpg" /> 
Figure 1: Density gradient medium-based separation of granulocytes. (A) Before and (B) after centrifugation. Note the sharp interface between the blood and the density gradient medium layers. (C) Sedimentation of the pellet in B resuspended (1:1) in 5% FBS/HBSS and 3% Dextran-0.9% NaCl. (D) PMN pellet after lysis of the residual RBCs in the supernatant of (C) with H2O. Abbreviations: PBMC = peripheral blood mononuclear cell; PMN = neutrophil; RBC = red blood cell; FBS = fetal bovine serum; HBSS = Hank&#39;s Balanced Salt Solution. <a href="https://www.jove.com/files/ftp_upload/62837/62837fig01large.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/62837/62837fig02.jpg" /> 
Figure 2: Spinning at lower speeds after RBC lysis decreased platelet contamination. After lysis of RBCs with H2O, cells were spun down at 200 &#215; g (A) or 400 &#215; g (B). Neutrophils were stained, and flow cytometry analysis was performed, as described in Figure 3, with the addition of anti-CD41 to label the platelets. Abbreviations: RBC = red blood cell; CD41 = cluster of differentiation 41; SSC-A = side scatter-Area. <a href="https://www.jove.com/files/ftp_upload/62837/62837fig02large.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/62837/62837fig03.jpg" /> 
Figure 3: Assessment of neutrophils isolated from buffy coat. Isolated neutrophils were stained with anti-CD66b, anti-CD11b, anti-CD14, and anti-CD193 following a standard protocol. Single cells (B) were gated from total cells (A). (C) CD11b+ cells were gated from single cells. (D) Dot plot showing neutrophils (CD66b+, CD14 low/-) and a very low monocyte contamination (CD66b-, CD14+, Q1). (E) CD66b- and CD66b+ cells were gated. (F) CD66b- cells were positive for lymphocyte markers (CD3, CD19). (G) Expression of CD11b and CD193 in CD66b+ cells, showing distinct neutrophil (CD66b+, CD11b+, CD193-) and eosinophil (CD66b+, CD11b-, CD193+) populations. In the representative result shown here, neutrophil purity is ~93% with ~3.7% lymphocyte and ~3.7% eosinophil contamination. (H) Quantification of neutrophil purity after purification. Data are compiled from 5 individual trials and presented as mean &#177; SD. Abbreviations: CD = cluster of differentiation; SSC-A = SSC-A = side scatter-Area; FSC-A = forward scatter-Area. <a href="https://www.jove.com/files/ftp_upload/62837/62837fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62837/62837fig04.jpg" /> 
Figure 4: Gradient purification did not cause CD62L shedding. (A-B) Control (A) or fMLP-stimulated neutrophils (B) (1 mM fMLP for 15 min at 37 &#176;C) were stained with neutrophil markers and CD62L. (C) Fluorescent mean intensity for CD62L is decreased in fMLP-treated cells, indicating CD62L shedding and neutrophil activation. Abbreviations: CD26L = L-selectin; fMLP = N-Formylmethionyl-leucyl-phenylalanine; SSC-A = SSC-A = side scatter-Area; FSC-A = forward scatter-Area; PE = phycoerythrin. <a href="https://www.jove.com/files/ftp_upload/62837/62837fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62837/62837fig05.jpg" /> 
Figure 5: Spontaneous death of neutrophils purified by density gradient or neutrophil isolation kit. Neutrophils were purified with density gradient (A and B) or commercial microbeads (C and D) and cultured for 0 h (A and C) or 24 h (B and D) in RPMI-10% FCS. Cells were stained using Annexin V and PI at respective time points following a standard protocol. (E) Quantification of purified neutrophil spontaneous death. n=5, mean &#177; SD. Abbreviations: SSC-A = SSC-A = side scatter-Area; FSC-A = forward scatter-Area; PI = propidium iodide; FCS = fetal calf serum; FITC = fluorescein isothiocyanate; AV5 = Annexin V. <a href="https://www.jove.com/files/ftp_upload/62837/62837fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation of Human Neutrophils from Whole Blood and Buffy Coats at JoVE.com

Isolation of Human Neutrophils from Whole Blood and Buffy Coats

This protocol details a method for the isolation of neutrophils from whole blood, buffy coats, or leukapheresis membranes, achieving good yield, high purity, and minimal cell activation. We utilize gradient purification, red blood cell (RBC) sedimentation, and RBC lysis to obtain a high-quality/purity neutrophil preparation.

Neutrophils (PMNs) are the most abundant leukocytes in human circulation, ranging from 40 to 70% of total blood leukocytes. They are the first cells ...

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17.7K Views.  Harvard Medical School, Department of Lab Medicine, Children’s Hospital Boston, and Dana-Farber/Harvard Cancer Center. This protocol details a method for the isolation of neutrophils from whole blood, buffy coats, or leukapheresis membranes, achieving good yield, high purity, and minimal cell activation. We utilize gradient purification, red blood cell (RBC) sedimentation, and RBC lysis to obtain a high-quality/purity neutrophil preparation.

Video: Isolation of Human Neutrophils from Whole Blood and Buffy Coats

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 ...

Isolation and Culture of Adult Epithelial Stem Cells from Human Skin

The homeostasis of all self-renewing tissues is dependent on adult stem cells. As undifferentiated stem cells undergo asymmetric divisions, they generate daughter cells that retain the stem cell phenotype and transit-amplifying cells (TA cells) that migrate from the stem cell niche, undergo rapid proliferation and terminally differentiate to repopulate the tissue.
Epithelial stem cells have been identified in the epidermis, hair follicle, and intestine as cells with a high in vitro proliferative potential and as slow-cycling label-retaining cells in vivo 1-3. Adult, tissue-specific stem cells are responsible for the regeneration of the tissues in which they reside during normal physiologic turnover as well as during times of stress 4-5. Moreover, stem cells are generally considered to be multi-potent, possessing the capacity to give rise to multiple cell types within the tissue 6. For example, rodent hair follicle stem cells can generate epidermis, sebaceous glands, and hair follicles 7-9. We have shown that stem cells from the human hair follicle bulge region exhibit multi-potentiality 10.
Stem cells have become a valuable tool in biomedical research, due to their utility as an in vitro system for studying developmental biology, differentiation, tumorigenesis and for their possible therapeutic utility. It is likely that adult epithelial stem cells will be useful in the treatment of diseases such as ectodermal dysplasias, monilethrix, Netherton syndrome, Menkes disease, hereditary epidermolysis bullosa and alopecias 11-13. Additionally, other skin problems such as burn wounds, chronic wounds and ulcers will benefit from stem cell related therapies 14,15. Given the potential for reprogramming of adult cells into a pluripotent state (iPS cells)16,17, the readily accessible and expandable adult stem cells in human skin may provide a valuable source of cells for induction and downstream therapy for a wide range of disease including diabetes and Parkinson's disease.

A rapid, robust way of isolating viable adult epithelial stem cells from human skin is described. The method utilizes enzymatic digestion of skin collagen matrix , followed by plucking of hair follicles and isolation of single cell suspensions or tissue fragments for cell culture.

The homeostasis of all self-renewing tissues is dependent on adult stem cells. As undifferentiated stem cells undergo asymmetric divisions, they generate ...

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 Primary Myofibroblasts from Mouse and Human Colon Tissue

The myofibroblast is a stromal cell of the gastrointestinal (GI) tract that has been gaining considerable attention for its critical role in many GI functions. While several myofibroblast cell lines are commercially available to study these cells in vitro, research results from a cell line exposed to experimental cell culture conditions have inherent limitations due to the overly reductionist nature of the work. Use of primary myofibroblasts offers a great advantage in terms of confirming experimental findings identified in a cell line. Isolation of primary myofibroblasts from an animal model allows for the study of myofibroblasts under conditions that more closely mimic the disease state being studied. Isolation of primary myofibroblasts from human colon tissue provides arguably the most relevant experimental data, since the cells come directly from patients with the underlying disease. We describe a well-established technique that can be utilized to isolate primary myofibroblasts from both mouse and human colon tissue. These isolated cells have been characterized to be alpha-smooth muscle actin and vimentin-positive, and desmin-negative, consistent with subepithelial intestinal myofibroblasts. Primary myofibroblast cells can be grown in cell culture and used for experimental purposes over a limited number of passages.

The myofibroblast is an influential stromal cell of the gastrointestinal tract that regulates important physiologic processes in both normal and disease states. We describe a technique that allows for the isolation of primary myofibroblasts from both mouse and human colon tissue, which can be utilized for in vitro experimentation.

The myofibroblast is  a stromal cell of the gastrointestinal (GI) tract that has been gaining  considerable attention for its critical role in many GI ...

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 ...

Isolation of Small Noncoding RNAs from Human Serum

The analysis of RNA and its expression is a common feature in many laboratories. Of significance is the emergence of small RNAs like microRNAs, which are found in mammalian cells. These small RNAs are potent gene regulators controlling vital pathways such as growth, development and death and much interest has been directed at their expression in bodily fluids. This is due to their dysregulation in human diseases such as cancer and their potential application as serum biomarkers. However, the analysis of miRNA expression in serum may be problematic. In most cases the amount of serum is limiting and serum contains low amounts of total RNA, of which small RNAs only constitute 0.4-0.5%1. Thus the isolation of sufficient amounts of quality RNA from serum is a major challenge to researchers today. In this technical paper, we demonstrate a method which uses only 400 &#181;l of human serum to obtain sufficient RNA for either DNA arrays or qPCR analysis. The advantages of this method are its simplicity and ability to yield high quality RNA. It requires no specialized columns for purification of small RNAs and utilizes general reagents and hardware found in common laboratories. Our method utilizes a Phase Lock Gel to eliminate phenol contamination while at the same time yielding high quality RNA. We also introduce an additional step to further remove all contaminants during the isolation step. This protocol is very effective in isolating yields of total RNA of up to 100 ng/&#181;l from serum but can also be adapted for other biological tissues.

This protocol describes a method for extracting small RNAs from human serum. We have used this method to isolate microRNAs from cancer serum for use in DNA arrays and also singleplex quantitative PCR. The protocol utilizes phenol and guanidinium thiocyanate reagents with modifications to yield high quality RNA.

The analysis of RNA and its expression is a common feature in many laboratories. Of significance is the emergence of small RNAs like microRNAs, which are ...

Isolation of Myofibroblasts from Mouse and Human Esophagus

Murine and human esophageal myofibroblasts are generated via enzymatic digestion. Neonate (8-12 day old) murine esophagus is harvested, minced, washed, and subjected to enzymatic digestion with collagenase and dispase for 25 min. Human esophageal resection specimens are stripped of muscularis propria and adventitia and the remaining mucosa is minced, and subjected to enzymatic digestion with collagenase and dispase for up to 6 hr. Cultured cells express &#945;-SMA and vimentin and express desmin weakly or not at all. Culture conditions are not conducive to growth of epithelial, hematopoietic, or endothelial cells. Culture purity is further confirmed by flow cytometric evaluation of cell surface marker expression of potential contaminating hematopoietic and endothelial cells. The described technique is straightforward and results in consistent generation of non-hematopoieitc, non-endothelial stromal cells. Limitations of this technique are inherent to the use of primary cultures in molecular biology studies, i.e., the unavoidable variability encountered among cultures established across different mice or humans. Primary cultures however are a more representative reflection of the in vivo state compared to cell lines. These methods also provide investigators the ability to isolate and culture stromal cells from different clinical and experimental conditions, allowing comparisons between groups. Characterized esophageal stromal cells can also be used in functional studies investigating epithelial-stromal interactions in esophageal disorders.

We present a protocol to generate primary cultures of murine and human esophageal stromal cells with a myofibroblast phenotype. Cultured cells have spindle shaped morphology, express &#945;-SMA and vimentin, and lack epithelial, hematopoietic and endothelial cell surface markers. Characterized stromal cells can be used in functional studies of epithelial-stromal interactions.

Murine and human esophageal myofibroblasts are generated via enzymatic digestion. Neonate (8-12 day old) murine esophagus is harvested, minced, washed, ...

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 Characterization of Microvesicles from Peripheral Blood

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived microvesicles (MVs, diameter &#62; 100 nm) is a fundamental cellular process that occurs in all living cells. These vesicles transport proteins, lipids and nucleic acids specific for their cell of origin and in vitro studies have highlighted their importance as mediators of intercellular communication. EVs have been successfully isolated from various body fluids and especially EVs in blood have been identified as promising biomarkers for cancer or infectious diseases. In order to allow the study of MV subpopulations in blood, we present a protocol for the standardized isolation and characterization of MVs from peripheral blood samples. MVs are pelleted from EDTA-anticoagulated plasma samples by differential centrifugation and typically possess a diameter of 100 - 600 nm. Due to their larger size, they can easily be studied by flow cytometry, a technique that is routinely used in clinical diagnostics and available in most laboratories. Several examples for quality control assays of the isolated MVs will be given and markers that can be used for the discrimination of different MV subpopulations in blood will be presented.

Extracellular vesicles present in blood have been suggested as novel biomarkers for various diseases. Here, we present a protocol for the isolation of large plasma membrane-derived microvesicles from peripheral blood samples and their subsequent analysis by conventional flow cytometry and Western Blotting.

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived ...

Nuclei Isolation from Whole Tissue using a Detergent and Enzyme-Free Method

High-throughput transcriptome and epigenome profiling requires preparation of a single cell or single nuclei suspension. Preparation of the suspension with&#160;intact cell or nuclei involves dissociation and permeabilization, steps that can introduce unwanted noise and undesirable damage. Particularly, certain cell-types such as neurons are challenging to dissociate into individual cells. Additionally, permeabilization of the cellular membrane to release nuclei requires optimization by trial-and-error, which can be time consuming, labor intensive and financially nonviable. To enhance the robustness and reproducibility of sample preparation for high-throughput sequencing, we describe a rapid enzyme and detergent-free column-based nuclei isolation method. The protocol enables efficient isolation of nuclei from the entire zebrafish brain within 20 minutes. The isolated nuclei display intact nuclear morphology and low propensity to aggregate. Further, flow cytometry allows nuclei enrichment and clearance of cellular debris for downstream application. The protocol, which should work on soft tissues and cultured cells, provides a simple and accessible method for sample preparation that can be utilized for high-throughput profiling, simplifying the steps required for successful single-nuclei RNA-seq and ATAC-seq experiments.

Single nuclei isolation relies on dissociation and detergent-based permeabilization of the cell membrane, steps that need optimization and are prone to introducing technical artifacts. We demonstrate a detergent and enzyme-free protocol for rapid isolation of intact nuclei directly from whole tissue, yielding nuclei suitable for single-nucleus RNA-seq (snRNA-Seq) or ATAC-seq.

High-throughput transcriptome and epigenome profiling requires preparation of a single cell or single nuclei suspension. Preparation of the suspension ...