Research has been supported by the Wallace H. Coulter Foundation Early Career Award in Translational Research. Thanks to Tim Andrews, Pediatric Hematology/Oncology, for help with blood sample collection. Also thanks to Dr. Sam Wellhausen, Flow Cytometry Core, James Graham Brown Cancer Center, for help with running flow cytometry samples.

Part 1. Microfluidic device fabrication
A silicon master mold of the microfluidic lysis device was created using standard photolithographic techniques with the equipment in the University of Louisville cleanroom [1]. AutoCAD (Autodesk, Inc., San Rafael, CA) was used to generate a transparency mask (Fineline Imaging, Colorado Springs, CO) to create negative replicas of the channels. The microfluidic lysis device was fabricated from the silicon master mold using soft lithographic techniques with the elastomer poly(dimethylsiloxane) (PDMS; Dow Corning, Midland, MI). The elastomer with the replicated channels was released, and channel access holes were punched with a 20-gauge needle (Small Parts, Miramar, FL.). The PDMS wafer was irreversibly bonded to a glass slide via oxygen plasma and tested for experimentation. 
Part 2. Protocol for running lysis device
After the microfluidic device has been fabricated and tested, experiments with erythrocyte lysis are ready to begin. The following section details the protocol for the microfluidic device.
2.1 Sample Collection
<ol>
 <li>Collect 4 milliliters of blood sample from median cubital vein, on the anterior forearm vein of patient with heparin as anticoagulant in two 2 ml green top vaccutainers.</li>
 <li>Discard first 2 ml tube as it contains dislodged mature endothelial cells (false positives).</li>
 <li>Resuspend the second tube to mix heparin with blood by turning up and down; save immediately on ice.</li>
 <li>Transfer 1 ml of blood to a 1.5 ml Eppendorf tube. Process the blood sample immediately (within one hour of collection).</li>
</ol>
2.2 Prime microfluidic device with PBS
<ol>
 <li>Obtain a microfluidic device bonded to glass. Press-fit access tubing (Small Parts, Miramar, FL) of slightly larger diameter in the inlets and outlet (Figure 1).</li>
 <li>Attach a 30-gauge syringe needle (Small Parts, Miramar, FL) to the tubing on each of the inlets, providing the macro to micro interface.</li>
 <li>Fill a 1 ml syringe with 1X PBS and make sure to get rid of bubbles by flicking the syringe needle. Connect the PBS-loaded 1 ml syringe to inlet 1 on the microfluidic device. Push the syringe containing the 1X PBS gently by hand until the solution flows out of inlet 2, and then immediately clamp inlet 2 with a standard office binder clip</li>
 <li>Continue pushing the 1X PBS until fluid reaches the outlet port of the microfluidics device. Once the solution flows out of the outlet, clamp the outlet tubing with another binder clip.</li>
 <li>Continue pushing the 1 ml syringe until the 1X PBS flows out of inlet 3, and clamp the inlet 3 port with another office binder clip. The microfluidics device is now fully primed.</li>
</ol>
<img src="/files/ftp_upload/1656/1656fig1.jpg" alt="Figure 1" /> Figure 1. Schematic of device showing inlets for respective solutions and outlet.
2.3 Preparation of syringe pumps and solution
<ol>
 <li>To calibrate the pumps follow these steps:
 <ol class="lalpha">
 <li>For the large Harvard syringe pump (Harvard, PHD 22/2000 Part # 702000): Set the diameter configuration to 22.5 mm, flow rate 600 &mu;l/min. </li>
 <li>For the small Harvard Syringe pump (Harvard, Pico Plus Part # 702213): Set the diameter configuration to 4.61 mm, flow rate 20 &mu;l/min. </li>
 </ol>
 </li>
 <li>Fill one 30 ml syringe with sterile deionized water. This can be accomplished by withdrawing the solution directly from the 50 ml conical tube. (Label the syringe)</li>
 <li>Fill a second 30 ml syringe with 2X phosphate-buffered-saline (PBS), 2% paraformaldehyde (PFA) using the procedure outlined in step 9. (Label the syringe)</li>
 <li>Fill a 1 ml syringe with 1X PBS from the aliquots stored in the 15 ml conical tubes, using the procedure outlined in step 9.</li>
 <li>Connect the 30 ml deionized water syringe to inlet 2 and the 30 ml 2X PBS syringe to inlet 3 (make sure you don t trap bubbles in the tubing or microfluidics device). Remove the clamp from inlets 2 and 3 and the outlet tubing. Connect the 1X PBS 1 ml syringe into inlet 1. Avoid introducing bubbles by filling the syringe needle with liquid, connecting the syringe, and flicking the syringe needle to rid of bubbles.</li>
</ol>
2.4 Erythrocyte lysis
<ol>
 <li>Set all of the syringes on the proper Harvard pumps as outlined below:
 <ol class="lalpha">
 <li>Set the two 30 ml syringes (one containing sterile, de-ionized water and the other with 2X PBS, 2% PFA) together on the large Harvard syringe pump.</li>
 <li>Set the 1 ml syringe (containing 1X PBS) on the small Harvard syringe pump. </li>
 <li>Place the outlet tubing in a waste collector (50 ml conical tube that is labeled waste ). </li>
 </ol>
 </li>
 <li>Turn on the large Harvard syringe pump (with the 30 ml syringes) and let it run for one minute. (Make sure there are no bubbles and no leakage in the device or in the tubing).</li>
 <li>Turn on the small Harvard syringe pump and let it run for an additional minute. (Make sure there are no bubbles and no leakage in the device or in the tubing). Stop the pumps. The microfluidics device is now ready to be used.</li>
 <li>Remove the 1 ml syringe containing 1X PBS from the small Harvard syringe pump.</li>
 <li>After obtaining a blood sample, fill the 1 ml syringe with 0.05 ml of sterile 1X PBS without trapping any bubbles. Next, fill the syringe with 0.5 ml of blood obtained from the Eppendorf tube. (The 1 ml syringe will now contain a final volume of 0.55 ml of the blood and 1X PBS buffer.) Note: Keep the syringe vertical while filling to avoid mixing of the 0.05 ml PBS with the blood. </li>
 <li>Connect the syringe containing the blood to inlet 1 and mount the syringe carefully (avoid pushing the blood through the tubing into the device) on the small Harvard syringe pump (the syringe is mounted vertical into the pump). </li>
 <li>Remove the outlet tubing from the waste tube and put a clean 50 ml centrifuge tube in its place and set it on a bucket of ice. </li>
 <li>Switch on the large Harvard syringe pump first and let it run for 1 minute. Start collecting the sample from the outlet into the 50 ml centrifuge tube. </li>
 <li>Switch on the small Harvard syringe pump.</li>
 <li>Once the blood sample in the 1 ml syringe has completely traveled through the device, stop both pumps. This should take approximately 20 minutes.</li>
 <li>Centrifuge the collected sample for 5 minutes at 350 x g at room temperature with the brake off.</li>
 <li>Remove supernatant by placing pipette tip at the opposite side of the white pellet. Remove as much supernatant as possible, especially red cell debris, without disturbing the white pellet.</li>
 <li>Resuspend the sample in 1 ml of flow buffer (1X PBS, 1% BSA [bovine serum albumin], 2% PFA) by pipetting up and down. Transfer sample to a 1.5 ml Eppendorf tube.</li>
 <li>Microcentrifuge the collected sample for 5 minutes at 4 x g at room temperature. Use same precision as before to remove the supernatant.</li>
 <li>Resuspend the sample in 1 ml of flow buffer by vortexing.</li>
</ol>
2.5 Preparation for flow cytometric analysis
<ol>
 <li>For sample analysis using flow cytometry, add 100 &mu;l of sample to a flow cytometry tube.</li>
 <li>To the 100 &mu;l of sample add specified antibody (ensure correct concentration).</li>
 <li>Allow sample to incubate for 30 minutes at 40 C. </li>
 <li>Wash twice with flow buffer prior to flow cytometry. Wash step includes adding 250 &mu;l of flow buffer, vortexing, centrifuging for 5 minutes at 350 x g, and removing supernatant with one fluid turn of flow cytometry tube. </li>
 <li>Resuspend the pellet in 250 &mu;l of flow buffer. Sample is ready for analysis by flow cytometry.</li>
</ol>
Representative Results
Whole blood has been run through the microfluidic lysis device and post-processing protocol, ridding of erythrocytes and enriching circulating nucleated cells (CNCs). Ideal results will yield a clean flow cytometry scatter plot with separate CNC populations. Displayed below in Figures 2 and 3 are scatter plot comparisons of a whole blood sample without erythrocyte lysis and with erythrocyte lysis by the microfluidic protocol with axes as forward scatter (FSC) versus side scatter (SSC) light. It can be seen that the microfluidic enrichment protocol is necessary to obtain a clean flow cytometry scatter plot. 
<img src="/files/ftp_upload/1656/1656fig2.jpg" alt="Figure 2" /> Figure 2. Flow cytometry scatter plot of whole blood sample without erythrocyte lysis with FSC and SSC axes.
<img src="/files/ftp_upload/1656/1656fig3.jpg" alt="Figure 3" /> Figure 3. Flow cytometry scatter plot of circulating nucleated cells with microfluidic erythrocyte lysis labeled by populations with FSC and SSC axes. Region 1 (red) represents lymphocytes, Region 2 (green) represents monocytes, Region 3 (blue) represents granulocytes, and Region 4 (purple) has yet to be characterized.
Removal of red cell debris by pipette in the protocol is also important when necessary. Below in Figure 4 is a scatter plot showing a surplus of red cell debris. Although not clouding the scatter plot as in Figure 2, one must account for the added events due to erythrocyte debris during data analysis.
<img src="/files/ftp_upload/1656/1656fig4.jpg" alt="Figure 4" /> Figure 4. Flow cytometry scatter plot depicting red cell debris, seen as a cluster of events in Region 5 (cyan).
Staining cells for certain antibody phenotype markers will also generate characteristic results. In the following Figures are antibody phenotype stains for 3 distinct leukocyte populations: lymphocytes, monocytes, and granulocytes. Figure 5 depicts lymphocyte populations plotted with axes CD3 and CD4. Monocytes and granulocytes are shown in Figures 6-7 with axes as FSC versus CD14 and CD66b, respectively.
<img src="/files/ftp_upload/1656/1656fig5.jpg" alt="Figure 5" /> Figure 5. Flow cytometry scatter plot depicting lymphocytes with CD3 and CD4 axes.
<img src="/files/ftp_upload/1656/1656fig6.jpg" alt="Figure 6" /> Figure 6. Flow cytometry scatter plot depicting monocytes with FSC and CD14 axes.
<img src="/files/ftp_upload/1656/1656fig7.jpg" alt="Figure 7" /> Figure 7. Flow cytometry scatter plot depicting granulocytes with FSC and CD66b axes.

<ol>
	<li>White, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical+application+of+microfluidic+leukocyte+enrichment+protocol+in+mild+phenotype+sickle+cell+disease+(SCD.">Clinical application of microfluidic leukocyte enrichment protocol in mild phenotype sickle cell disease (SCD.</a> Biomedical Microdevices. 11 (2), 477-483 (2009).</li><li>Sethu, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continuous+Flow+Microfluidic+Device+for+Rapid+Erythrocyte+Lysis.">Continuous Flow Microfluidic Device for Rapid Erythrocyte Lysis.</a> Anal. Chem. 76 (21), 6247-6253 (2004).</li></ol>

An automated microfluidic blood lysis protocol has been reported. Within the protocol are critical steps worth noting. In section P.2.1, it is important to discard the first blood vial collected as the sample contains dislodged mature endothelial cells acting as false positives. Also make sure to run the blood sample within an hour of collection. Priming the microfluidic device in section P.2.2, it is important to initially rid of the bubbles. Additionally, one should avoid introducing bubbles when attaching new syringes...

automated microfluidic blood lysis protocol for enrichment of circulating nucleated cells

In this report the protocol for an automated microfluidic blood lysis device is detailed. Circulating nucleated cells (CNCs), including leukocytes and endothelial cells, provide an ideal platform for an updated status on the immune condition of an individual. The microfluidic protocol allows for enrichment of CNCs without selective cell loss and sample preparation variability due to user-mediated steps. Briefly, the protocol includes device fabrication, sample collection, device setup, and running blood through the microfluidic chamber. Within the device whole blood is rapidly mixed with deionized water for approximately 10 seconds in a 50 micron x 150 micron microfluidic channel. In this time span erythrocytes are lysed due to hypotonic conditions. Herringbone structures on the bottom of the channel ensure thorough mixing and exposure of cells to a constant environment. Remaining cells are returned to isotonic conditions at the exit of the device, fixed using 2% paraformaldehyde, centrifuged to separate erythrocyte debris from CNCs, and suspended in flow buffer for staining and analysis by flow cytometry. Results show clean flow cytometry scatter plots with CNC populations saved. Significance of this device and protocol comes in the study and understanding of disease pathogenesis by analysis of CNC populations. Hence, automation, effectiveness, and simplicity of the microfluidic protocol are demonstrated.

Watch this Scientific Journal Video about Automated Microfluidic Blood Lysis Protocol for Enrichment of Circulating Nucleated Cells at JoVE.com

Automated Microfluidic Blood Lysis Protocol for Enrichment of Circulating Nucleated Cells

An automated microfluidic device was developed for circulating nucleated cell enrichment from peripheral blood via erythrocyte lysis that ensures isolation of high quality sample without cell loss.

In this report the protocol for an automated microfluidic blood lysis device is detailed. Circulating nucleated cells (CNCs), including leukocytes and ...

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12.3K Views.  University of Louisville. An automated microfluidic device was developed for circulating nucleated cell enrichment from peripheral blood via erythrocyte lysis that ensures isolation of high quality sample without cell loss.

Video: Automated Microfluidic Blood Lysis Protocol for Enrichment of Circulating Nucleated Cells

Enrichment of NK Cells from Human Blood with the RosetteSep Kit from StemCell Technologies

Natural killer (NK) cells are large granular cytotoxic lymphocytes that belong to the innate immune system and play major roles in fighting against cancer and infections, but are also implicated in the early stages of pregnancy and transplant rejection. These cells are present in peripheral blood, from which they can be isolated. Cells can be isolated using either positive or negative selection. For positive selection we use antibodies directed to a surface marker present only on the cells of interest whereas for negative selection we use cocktails of antibodies targeted to surface markers present on all cells but the cells of interest. This latter technique presents the advantage of leaving the cells of interest free of antibodies, thereby reducing the risk of unwanted cell activation or differenciation. In this video-protocol we demonstrate how to separate NK cells from human blood by negative selection, using the RosetteSep kit from StemCell technologies. The procedure involves obtaining human peripheral blood (under an institutional review board-approved protocol to protect the human subjects) and mixing it with a cocktail of antibodies that will bind to markers absent on NK cells, but present on all other mononuclear cells present in peripheral blood (e.g., T lymphocytes, monocytes...). The antibodies present in the cocktail are conjugated to antibodies directed to glycophorin A on erythrocytes. All unwanted cells and red blood cells will therefore be trapped in complexes. The mix of blood and antibody cocktail is then diluted, overlayed on a Histopaque gradient, and centrifuged. NK cells (&gt;80% pure) can be collected at the interface between the Histopaque and the diluted plasma. Similar cocktails are available for enrichment of other cell populations, such as human T lymphocytes.

Natural killer cells are a small population of lymphocytes. Here we show how to isolate these cells from human blood by negative selection, using a kit from StemCell Technologies. The cells obtained are viable and untouched by antibodies, and therefore ready to be used for a number of procedures.

Natural killer (NK) cells are large granular cytotoxic lymphocytes that belong to the innate immune system and play major roles in fighting against cancer ...

Microfluidic Co-culture of Epithelial Cells and Bacteria for Investigating Soluble Signal-mediated Interactions

The human gastrointestinal (GI) tract is a unique environment in which intestinal epithelial cells and non-pathogenic (commensal) bacteria coexist. It has been proposed that the microenvironment that the pathogen encounters in the commensal layer is important in determining the extent of colonization. Current culture methods for investigating pathogen colonization are not well suited for investigating this hypothesis as they do not enable co-culture of bacteria and epithelial cells in a manner that mimics the GI tract microenvironment. Here we describe a microfluidic co-culture model that enables independent culture of eukaryotic cells and bacteria, and testing the effect of the commensal microenvironment on pathogen colonization. The co-culture model is demonstrated by developing a commensal Escherichia coli biofilm among HeLa cells, followed by introduction of enterohemorrhagic E. coli (EHEC) into the commensal island, in a sequence that mimics the sequence of events in GI tract infection.

This protocol describes a microfluidic co-culture model for simultaneous and localized culture of epithelial cells and bacteria. This model can be used for investigating the role of different soluble molecular signals on pathogenesis as well as screen the effectiveness of putative probiotic bacterial strains.

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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 ...

Immunofluorescence Analysis of Endogenous and Exogenous Centromere-kinetochore Proteins

"Centromeres" and "kinetochores" refer to the site where chromosomes associate with the spindle during cell division. Direct visualization of centromere-kinetochore proteins during the cell cycle remains a fundamental tool in investigating the mechanism(s) of these proteins. Advanced imaging methods in fluorescence microscopy provide remarkable resolution of centromere-kinetochore components and allow direct observation of specific molecular components of the centromeres and kinetochores. In addition, methods of indirect immunofluorescent (IIF) staining using specific antibodies are crucial to these observations. However, despite numerous reports about IIF protocols, few discussed in detail problems of specific centromere-kinetochore proteins.1-4 Here we report optimized protocols to stain endogenous centromere-kinetochore proteins in human cells by using paraformaldehyde fixation and IIF staining. Furthermore, we report protocols to detect Flag-tagged exogenous CENP-A proteins in human cells subjected to acetone or methanol fixation. These methods are useful in detecting and quantifying endogenous centromere-kinetochore proteins and Flag-tagged CENP-A proteins, including those in human cells.

Here we report protocols to detect endogenous and exogenous centromere-kinetochore proteins in human cells and quantify these protein levels at centromeres-kinetochores by indirect immunofluorescent staining through the use of fixation (paraformaldehyde, acetone, or methanol fixation).

"Centromeres" and "kinetochores" refer to the site where chromosomes associate with the spindle during cell division. Direct visualization of ...

Using Microfluidic Devices to Measure Lifespan and Cellular Phenotypes in Single Budding Yeast Cells

Budding yeast Saccharomyces cerevisiae is an important model organism in aging research. Genetic studies have revealed many genes with conserved effects on the lifespan across species. However, the molecular causes of aging and death remain elusive. To gain a systematic understanding of the molecular mechanisms underlying yeast aging, we need high-throughput methods to measure lifespan and to quantify various cellular and molecular phenotypes in single cells. Previously, we developed microfluidic devices to track budding yeast mother cells throughout their lifespan while flushing away newborn daughter cells. This article presents a method for preparing microfluidic chips and for setting up microfluidic experiments. Multiple channels can be used to simultaneously track cells under different conditions or from different yeast strains. A typical setup can track hundreds of cells per channel and allow for high-resolution microscope imaging throughout the lifespan of the cells. Our method also allows detailed characterization of the lifespan, molecular markers, cell morphology, and the cell cycle dynamics of single cells. In addition, our microfluidic device is able to trap a significant amount of fresh mother cells that can be identified by downstream image analysis, making it possible to measure the lifespan with higher accuracy.

This article presents a protocol optimized for the production of microfluidic chips and the setup of microfluidic experiments to measure the lifespan and cellular phenotypes of single yeast cells.

Budding yeast Saccharomyces cerevisiae is an important model organism in aging research. Genetic studies have revealed many genes with conserved effects ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

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A Protocol for Using Gene Set Enrichment Analysis to Identify the Appropriate Animal Model for Translational Research

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We provide a standardized protocol for the use of gene set enrichment analysis of transcriptomic data to identify an ideal mouse model for translational research. 
This protocol can be used with DNA microarray and RNA sequencing data and can further be extended to other omics data if data are available.

Recent studies that compared transcriptomic datasets of human diseases with datasets from mouse models using traditional gene-to-gene comparison ...

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the lymph node microenvironment in B-cell non-Hodgkin&#39;s lymphoma is analyzed. Subsets of circulating T-follicular helper cells (cTfh), the circulating memory compartment of Tfh cells in the blood, are also perturbed in disease and therefore represent potential novel predictive biomarkers. Peripheral blood-based testing is advantageous because it is relatively non-invasive and allows simple serial monitoring.This article describes a method for isolating CD4+ T-cells from human blood, and further analysis by flow-cytometry to enumerate cTfh cells and the proportions of their various subsets (cTfhPD-1-/+/hi, cTfh1,2,17 and cTfh1/17). The level of these subsets was then compared between normal subjects and patients with lymphoma. We found that the method was robust enough to obtain reliable results from routinely collected patient material. The technique we describe for the analysis can be easily adapted to cell sorting and downstream applications such as RT-PCR.

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the ...

Isolation and Enrichment of Human Lung Epithelial Progenitor Cells for Organoid Culture

Epithelial organoid models serve as valuable tools to study the basic biology of an organ system and for disease modeling. When grown as organoids, epithelial progenitor cells can self-renew and generate differentiating progeny that exhibit cellular functions similar to those of their in vivo counterparts. Herein we describe a step-by-step protocol to isolate region-specific progenitors from human lung and generate 3D organoid cultures as an experimental and validation tool. We define proximal and distal regions of the lung with the goal of isolating region-specific progenitor cells. We utilized a combination of enzymatic and mechanical dissociation to isolate total cells from the lung and trachea. Specific progenitor cells were then fractionated from the proximal or distal origin cells using fluorescence associated cell sorting (FACS) based on cell type-specific surface markers, such as NGFR&#160;for sorting basal cells and HTII-280 for sorting alveolar type II cells. Isolated basal or alveolar type II progenitors were used to generate 3D organoid cultures. Both distal and proximal progenitors formed organoids with a colony forming efficiency of 9-13% in distal region and 7-10% in proximal region when plated 5000 cell/well on day 30. Distal organoids maintained HTII-280+ alveolar type II cells in culture whereas proximal organoids differentiated into ciliated and secretory cells by day 30. These 3D organoid cultures can be used as an experimental tool for studying the cell biology of lung epithelium and epithelial mesenchymal interactions, as well as for the development and validation of therapeutic strategies targeting epithelial dysfunction in a disease.

This article provides a detailed methodology for tissue dissociation and cellular fractionation approaches allowing enrichment of viable epithelial cells from proximal and distal regions of the human lung. Herein these approaches are applied for the functional analysis of lung epithelial progenitor cells through the use of 3D organoids culture models.

Epithelial organoid models serve as valuable tools to study the basic biology of an organ system and for disease modeling. When grown as organoids, ...

Quantification of Circulating Pig-Specific DNA in the Blood of a Xenotransplantation Model

Xenotransplantation is a feasible method to treat organ failure. However, how to effectively monitor the immune rejection of xenotransplantation is a problem for physicians and researchers. This manuscript describes a simple and effective method to monitor immune rejection in pig-to-mouse cell transplantation models and pig-to-monkey artery patch transplantation models. Circulating DNA is a potentially non-invasive biomarker for organ damage. In this study, circulating pig-specific DNA (cpsDNA) was monitored during xenograft rejection by quantitative real-time PCR (qPCR). In this protocol, porcine specific primers were designed, plasmids-containing porcine specific DNA fragments were constructed, and standard curves for quantitation were established. Species-specific primers were then used to quantify cpsDNA by qPCR in pig-to-mouse cell transplantation models and pig-to-monkey artery patch transplantation models. The value of this method suggests that it can be used as a simple, convenient, low cost, and less invasive method to monitor the immune rejection of xenotransplantation.

In this protocol, porcine specific primers were designed, plasmids-containing porcine specific DNA fragments were constructed, and standard curves for quantitation were established. Using species-specific primers, cpsDNA was quantified by qPCR in pig-to-mouse cell transplantation models and pig-to-monkey artery patch transplantation models.

Xenotransplantation is a feasible method to treat organ failure. However, how to effectively monitor the immune rejection of xenotransplantation is a ...