Name: purification of specific cell population by fluorescence activated cell sorting (facs)
Uploaded: 2010-07-10T18:30:00
Duration: 15 min 29 s
Description: Watch this Scientific Journal Video about Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS at JoVE.com

We would like to thank Dr. Bill Cashdollar for his support in the use of the Flow Cytometry Core.

<ol><li>Before starting the process, the following items need to be to collected and prepared:
<ol style="list-style-type:lower-roman;">
 <li>Ice</li>
 <li>15 ml conical tubes for staining cells and 12 x 75 mm flow tubes for staining single color controls.</li>
 <li>Staining buffer: phosphate buffered saline (PBS) + 3% fetal calf serum (FCS)</li>
 <li>Pipettes</li>
 <li>Fc block (if required) and staining antibodies. Antibodies need to be titrated for optimal staining.</li>
 <li>Compensation beads</li>
 <li>Suspension buffer: Hank's Balanced Salt Solution (HBSS) + 25 mM HEPES + 3% FCS</li>
 <li>Trypan blue and hemocytometer</li>
 <li>Cell strainer (40 &mu;m Nylon)</li>
 <li>Collection tubes: Two types of collection tubes can be used a) 12 x 75 mm polystyrene tubes (containing 300 &mu;l FCS and 25 mM HEPES) or b) 15 ml conical tubes (containing 1 ml FCS + 25 mM HEPES). Any rich medium with high serum concentration can be used for the collection of sorted cells. </li>
</ol></li>
<li>Generate a single cell suspension of the starting cell population.</li>
<li>Optional: Enrich for the desired cell population by a bulk purification method such as complement depletion or magnetic sorting. The main advantage of the enrichment step is that it reduces the sort time.</li>
<li>Wash cells once with staining buffer.</li>
<li>Discard the supernatant and resuspend the cells in staining buffer at a concentration up to 50 x 106 for efficient staining.</li>
<li>Optional: For cells expressing high levels of FcR, block the receptors using an appropriate blocking method. A method of choice is the use of a mAb that binds to Fc&gamma;R on ice for 10-15 min. These antibodies are commercially available.</li>
<li>Add the appropriate mAb (at a predetermined concentration) to stain the desired cell population and incubate for 20-30 min on ice in the dark, followed by two washes with staining buffer. Optional: Staining with a vital dye, such as propidium iodide, can be included to discern dead cells (5). If multi-color staining is to be used, single color controls are required. We use BD CompBeads to prepare single color controls using the manufacturer's protocol.</li>
<li>If not using directly conjugated antibodies, repeat step 7 using the appropriate secondary antibody or Streptavidin conjugate.</li>
<li>After washing, resuspend the cells in culture medium and determine the cell concentration using a vital dye such as Trypan blue.</li>
<li>Adjust the cell concentration to 15-20 x 106/ml. For cell populations that form clusters, which can clog the instrument during sorting, filter the cells through a strainer.</li>
<li>Set up and optimize the cell sorter. The process of setting up a flow cytometer is varied depending on the manufacture and needs to be performed by appropriately trained personnel. The general recommendations are as follows:
<ol style="list-style-type:lower-roman;">
 <li>Select the appropriate nozzle depending on the cell type to be sorted.</li>
 <li>For sterile sorts, sterilize the instrument.</li>
 <li>Perform instrument quality control with beads to verify lasers are functioning, and it is sorting accurately. </li>
 <li>Install the required collection device and set up the side streams.</li>
 <li>Look at the instrument's laser and detector set up to determine the fluorescent labels that can be used (our BD FACS Aria has three lasers and can detect up to nine colors).</li>
 <li>Once it is determined what fluorescent labels will be used on the cell sorter, compensation can be performed (as explained below).</li>
</ol></li>
<li>Perform compensation using the negative control sample and the single positive controls. Compensation is necessary to remove the spectrum overlap between two detectors. It is pertinent to note that compensation is not fool proof and is adversely affected by how brightly certain markers stain, and by the fluorescence of cells that have low autofluorescence which can lead to poor resolution between dim and negative populations. 
 For best results, the experimental design should include using bright fluorochromes with markers that have low expression or that don't have a known staining pattern. Markers that provide good separation between cell populations and are highly expressed should be used with dimer fluorochromes such as ones excited by red or violet lasers.
<ol style="list-style-type:lower-roman;">
 <li>Compensation can be performed with compensation beads, which are polystyrene microparticles that have been coupled to an antibody specific for the Kappa light chain of Ig from mouse, rat, or rat/hamster. The beads are easy to stain, have a robust signal and provide an easy way of preparing single stained controls that have the same fluorophore as the experimental samples.</li>
 <li>Set up a template that includes a bivariate plot to display forward scatter (FSC) and side scatter (SSC), and one histogram for each fluorochrome that will be used. </li>
 <li>Run the negative control tube and adjust the forward scatter (FSC) and side scatter (SSC) to place the population of interest on scale.</li>
 <li>Adjust the fluorescence PMT settings to place the negative population or autofluorescence in the far left hand portion of the histogram.</li>
 <li>Record the negative control tube.</li>
 <li>Run the single positive control tubes and record the data for each tube. </li>
 <li>Draw an interval gate around the positive portion of the data on the histogram, and another interval gate on the negative portion of the histogram.</li>
 <li>Manual compensation is done by adjusting the median (median is a better estimate of central tendency than the mean on a logarithmic scale) of the positives until it is equal to the median of the negatives. This must be done for each of the fluorochrome labels used in the experiment. </li>
 <li>To adjust the median, adjust the spectral overlap values either higher or lower until the medians for each fluorescent parameter match as closely as possible.</li>
 <li>Compensation can also be done automatically with analysis software. Automatic compensation uses the recorded data from compensation controls to calculate an algebraic matrix for all the fluorophores used in the experiment. These values are used to subtract out the contributions of the non-primary colors that are bleeding into a primary color's detector. </li>
</ol></li>
<li>Record the experimental sample to be sorted and use gating tools and subsetting methods to define the populations of interest.
<ol style="list-style-type:lower-roman;">
 <li>Set up the experimental template with a dot plot that displays FSC vs. SSC and use a gating tool, polygon, rectangle, interval or quadrant to gate the population of interest. The scatter plot displays the physical properties of the cell which is distinct to the cell type. </li>
 <li>The best way to determine the fluorescence gating strategy is to use fluorescence minus one (FMO) controls. In a FMO control tube all reagents that are used in the experiment are included except for the one of interest. The FMO helps discriminate between dimly stained populations and broad negative populations. </li>
 <li>Use the data recorded from FMO tubes to determine where to place gates within the experimental tube. </li>
</ol></li>
<li>Once gates have been determined, the gates can be selected for sorting into external collection tubes. Up to four gates (or populations) can be sorted at one time depending on the instrumentation available.</li>
<li>Run the experimental sample tube at 4 &deg;C, turn on deflection plates, and sort the sample. 5 x 105 -1.5 x 106 cells can be sorted into a 12 x 75 mm tube and 1.5 x 106 - 4.5 x 106 cells can be sorted into a 15 ml conical.</li>
<li>Once the required number of cells has been obtained, manually stop sorting.</li>
<li>Perform a post-sort analysis to determine the purity of the sorted cell populations.</li>
</ol>
Representative results
We have shown here results for mouse splenic B and CD4 T cell sorting (Figure 1). Splenocytes were stained with PE Texas Red-conjugated anti-mouse B220, FITC-conjugated anti-mouse TCR&beta; and Alexafluor-700-conjugated anti-mouse CD4 to identify B cells (B220+TCR&beta;-) and CD4 T cells (CD4+TCR&beta;+). The sorting was performed with BD FACS Aria instrument. After sorting the B cell purity was &gt;97% and the CD4 T cell purity was &gt;98%.
<img src="/files/ftp_upload/1546/1546fig1.jpg" alt="Figure 1" /> Figure 1. Purity of splenic B cells and CD4 T cells after FACS. Mouse splenocytes were stained with anti-mouse B220, TCR&beta; and CD4 antibodies. FACS for B cells was performed using a BD FACS Aria by gating on B220+TCR&beta;- cells (Fig. 1A, left panel, lower right quadrant). A post sort analysis was performed to determine the purity of the B cells (Fig. 1A, right panel). B220 staining is shown on the x-axis and TCR&beta; staining on the y-axis. FACS for CD4 T cells was performed using the same flow cytometer by gating on TCR&beta;+CD4+ cells (Fig. 1B, left panel, upper right quadrant). A post sort analysis was performed to determine the purity of the CD4 T cells (Fig. 1B, right panel). TCR&beta; staining is shown on the x-axis and CD4 staining on the y-axis. The percent positive cells in each quadrant is shown.

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FACS is a highly sophisticated technique for purifying cell populations of interest, in which a very high purity (95-100%) of the sorted population can be obtained. Therefore, this technique is particularly important for experiments, where high purity is an essential requirement (e.g. microarray analysis). FACS is particularly advantageous over other available methods of purification when a cell population needs to be purified based on a weekly expressed surface marker or when two or more populations need to be purified which have different levels of expression of the same surface marker. For example, the purification of marginal zone B cells (B220+CD21hiCD23int/low) and follicular B cells (B220+CD21int/lowCD23hi) based on expression levels of CD21 and CD23. Another growing application of FACS is single cell sorting allowing the analysis of individual cells (3, 4). FACS is also commonly used to sort cells expressing fluorescent proteins that are genetically expressed, such as green fluorescent protein tagged proteins (5). If a sterile sort is performed the cells can be cultured. However, cell viability and yield can be compromised during sorting. Viability can be improved by using a cell-dependent optimal temperature for the sorting and by processing collection tubes immediately after they reach capacity. Recovery can be increased by using polypropylene collection tubes, collecting sorted cells in serum rich media and inverting collection tubes intermittently, maintaining collection tubes at an optimal temperature and centrifuging sorted cells for ~10 min (http://cyto.mednet.ucla.edu/Protocol.htm).
The development of ultra high-speed sorters has extended the possibility of the application of flow sorting in clinical settings. The potential clinical applications of FACS include purification of blood stem cells from human blood for therapeutic purposes, applications in cancer therapy, amniocentesis replacement, sorting human sperm and early disease detections (6, 7).
For multicolor staining, antibody selection is a critical step and fluorochromes are chosen based on the detection system for each particular instrument. There are a number of companies that sell cell sorters and the laser configuration varies according to user needs. Thus one cannot simply use the exact same antibodies used in published studies without confirming compatibility. For optimal antibody selection, one should consider the expression level of the protein being examined. In general, the brightest available fluorochrome-tagged antibodies should be used to stain dimly expressed surface markers, while dim fluorochromes can be used for the staining of highly expressed surface markers (8). In addition, antibodies should be selected in such a way that overlap among their emission spectra is minimal. For multicolor staining, it is extremely difficult to choose fluorochrome that have no spectral overlap. In this situation, compensation is performed.
Compensation is necessary to mathematically eliminate the fluorescence overlap between the emissions spectrum of the various fluorochromes that is measureable by the detectors (8-10). In other words, the overlap fluorescence from one fluorochrome is subtracted from the emissions spectrum of another fluorochrome. The compensation calculation requires an unstained control and single positive controls for each fluorochrome used for a multicolor staining (9, 10). Compensation can be conducted using cells or beads. The beads are easy to stain, have a robust signal and provide an easy way of preparing single stained controls that have the same fluorophores as the experimental samples. Compensation beads are particularly advantageous when the cell surface protein of interest is expressed at a low level and when the cell population of interest is limiting in the initial starting cell population.
Once the antibodies have been chosen, their activity should be optimized by performing a titration analysis. This is particularly important if the FACS machine has highly powerful lasers. The use of suboptimal concentrations of staining antibodies can result in poor separation of the desired cell population from the total cell population. The use of high antibody concentrations increases the chance for antigen non-specific staining. Therefore antibody titration will allow the selection of the antibody concentration that gives the maximum brightness of the positive population and minimal background staining (8).
FACS is now a standard technique for the purification of subpopulations of cells. It can be used to separate any cell type in which a single cell suspension can be generated and antibodies are available to identify the desired cell population. FACS is the method of choice when highly pure cell populations are required.

purification of specific cell population by fluorescence activated cell sorting (facs)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known phenotype.  Other bulk methods of purification include panning, complement depletion and magnetic bead separation.  However, FACS has several advantages over other available methods.  FACS is the preferred method when very high purity of the desired population is required, when the target cell population expresses a very low level of the identifying marker or when cell populations require separation based on differential marker density.  In addition, FACS is the only available purification technique to isolate cells based on internal staining or intracellular protein expression, such as a genetically modified fluorescent protein marker.  FACS allows the purification of individual cells based on size, granularity and fluorescence.  In order to purify cells of interest, they are first stained with fluorescently-tagged monoclonal antibodies (mAb), which recognize specific surface markers on the desired cell population (1).  Negative selection of unstained cells is also possible.  FACS purification requires a flow cytometer with sorting capacity and the appropriate software.  For FACS, cells in suspension are passed as a stream in droplets with each containing a single cell in front of a laser.  The fluorescence detection system detects cells of interest based on predetermined fluorescent parameters of the cells.  The instrument applies a charge to the droplet containing a cell of interest and an electrostatic deflection system facilitates collection of the charged droplets into appropriate collection tubes (2).  The success of staining and thereby sorting depends largely on the selection of the identifying markers and the choice of mAb.  Sorting parameters can be adjusted depending on the requirement of purity and yield.  Although FACS requires specialized equipment and personnel training, it is the method of choice for isolation of highly purified cell populations.

Watch this Scientific Journal Video about Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS at JoVE.com

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity. Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known ...

purification-specific-cell-population-fluorescence-activated-cell

Research

JoVE Journal

Immunology and Infection

Depletion of Specific Cell Populations by Complement Depletion

The purification of immune cell populations is often required in order to study their unique functions. In particular, molecular approaches such as real-time PCR and microarray analysis require the isolation of cell populations with high purity. Commonly used purification strategies include fluorescent activated cell sorting (FACS), magnetic bead separation and complement depletion. Of the three strategies, complement depletion offers the advantages of being fast, inexpensive, gentle on the cells and a high cell yield. The complement system is composed of a large number of plasma proteins that when activated initiate a proteolytic cascade culminating in the formation of a membrane-attack complex that forms a pore on a cell surface resulting in cell death1. The classical pathway is activated by IgM and IgG antibodies and was first described as a mechanism for killing bacteria. With the generation of monoclonal antibodies (mAb), the complement cascade can be used to lyse any cell population in an antigen-specific manner. Depletion of cells by the complement cascade is achieved by the addition of complement fixing antigen-specific antibodies and rabbit complement to the starting cell population. The cells are incubated for one hour at 37&deg;C and the lysed cells are subsequently removed by two rounds of washing. MAb with a high efficiency for complement fixation typically deplete 95-100% of the targeted cell population. Depending on the purification strategy for the targeted cell population, complement depletion can be used for cell purification or for the enrichment of cell populations that then can be further purified by a subsequent method.

To effectively study the function of immune cell populations their purification is often required. Complement depletion is a fast and inexpensive technique for the isolation of immune cell populations with high purity.

The purification of immune cell populations is often required in order to study their unique functions.  In particular, molecular approaches such as ...

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method enables researchers to better understand the characteristics of a single cell population without the influence of other cells. Compared to other methods of cell enrichment, such as magnetic-activated cell sorting (MCS), FACS is more flexible and accurate for cell separation due to the ability of phenotype detection by flow cytometry. In addition, FACS is usually capable of separating multiple cell populations simultaneously, which improves the efficiency and diversity of experiments. Although FACS has some limitations, it has been broadly used to purify cells for functional studies in both in vitro and in vivo settings. Here we report a protocol using fluorescence-activated cell sorting to isolate a very rare population of immune cells, plasmacytoid dendritic cells (pDC), with high purity from the bone marrow of lupus-prone mice for in vitro functional studies of pDC.

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of pDC.

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method ...

Generating De Novo Antigen-specific Human T Cell Receptors by Retroviral Transduction of Centric Hemichain

T cell receptors (TCRs) are used clinically to direct the specificity of T cells to target tumors as a promising modality of immunotherapy. Therefore, cloning TCRs specific for various tumor-associated antigens has been the goal of many studies. To elicit an effective T cell response, the TCR must recognize the target antigen with optimal affinity. However, cloning such TCRs has been a challenge and many available TCRs possess sub-optimal affinity for the cognate antigen. In this protocol, we describe a method of cloning de novo high affinity antigen-specific TCRs using existing TCRs by exploiting hemichain centricity. It is known that for some TCRs, each TCR&#945; or TCR&#946; hemichain do not contribute equally to antigen recognition, and the dominant hemichain is referred to as the centric hemichain. We have shown that by pairing the centric hemichain with counter-chains differing from the original counter-chain, we are able to maintain the antigen specificity, while modulating its interaction strength for the cognate antigen. Thus, the therapeutic potential of a given TCR can be improved by optimizing the pairing between the centric and counter hemichains.

Generating De Novo Antigen-specific Human T Cell Receptors by Retroviral Transduction of Centric Hemichain

Herein we describe a novel method to generate antigen-specific T cell receptors (TCRs) by pairing the TCR&#945; or TCR&#946; of an existing TCR, possessing the antigen-specificity of interest, with complementary hemichain of the peripheral T cell receptor repertoire. The de novo generated TCRs retain antigen-specificity with varying affinity.

T cell receptors (TCRs) are used clinically to direct the specificity of T cells to target tumors as a promising modality of immunotherapy. Therefore, ...

Detection of Trypanosoma brucei Variant Surface Glycoprotein Switching by Magnetic Activated Cell Sorting and Flow Cytometry

Trypanosoma brucei, a protozoan parasite that causes both Human and Animal African Trypanosomiasis (known as sleeping sickness and nagana, respectively) cycles between a tsetse vector and a mammalian host. It evades the mammalian host immune system by periodically switching the dense, variant surface glycoprotein (VSG) that covers its surface. The detection of antigenic variation in Trypanosoma brucei can be both cumbersome and labor intensive. Here, we present a method for quantifying the number of parasites that have 'switched' to express a new VSG in a given population. The parasites are first stained with an antibody against the starting VSG, and then stained with a secondary antibody attached to a magnetic bead. Parasites expressing the starting VSG are then separated from the rest of the population by running the parasites over a column attached to a magnet. Parasites expressing the dominant, starting VSG are retained on the column, while the flow-through contains parasites that express a new VSG as well as some contaminants expressing the starting VSG. This flow-through population is stained again with a fluorescently labeled antibody against the starting VSG to label contaminants, and propidium iodide (PI), which labels dead cells. A known number of absolute counting beads that are visible by flow cytometry are added to the flow-through population. The ratio of beads to number of cells collected can then be used to extrapolate the number of cells in the entire sample. Flow cytometry is used to quantify the population of switchers by counting the number of PI negative cells that do not stain positively for the starting, dominant VSG. The proportion of switchers in the population can then be calculated using the flow cytometry data.

Detection of Trypanosoma brucei Variant Surface Glycoprotein Switching by Magnetic Activated Cell Sorting and Flow Cytometry

African trypanosomes grown in vitro undergo antigenic variation at a low rate, such that populations are made up of parasites expressing a dominant variant surface glycoprotein (VSG) type and a small population of "switched" variants. This protocol describes a fast method for detecting and quantifying these populations.

Trypanosoma brucei, a protozoan parasite that causes both Human and Animal African Trypanosomiasis (known as sleeping sickness and nagana, respectively) ...

Molecular Diffusion in Plasma Membranes of Primary Lymphocytes Measured by Fluorescence Correlation Spectroscopy

Fluorescence correlation spectroscopy (FCS) is a powerful technique for studying the diffusion of molecules within biological membranes with high spatial and temporal resolution. FCS can quantify the molecular concentration and diffusion coefficient of fluorescently labeled molecules in the cell membrane. This technique has the ability to explore the molecular diffusion characteristics of molecules in the plasma membrane of immune cells in steady state (i.e., without processes affecting the result during the actual measurement time). FCS is suitable for studying the diffusion of proteins that are expressed at levels typical for most endogenous proteins. Here, a straightforward and robust method to determine the diffusion rate of cell membrane proteins on primary lymphocytes is demonstrated. An effective way to perform measurements on antibody-stained live cells and commonly occurring observations after acquisition are described. The recent advancements in the development of photo-stable fluorescent dyes can be utilized by conjugating the antibodies of interest to appropriate dyes that do not bleach extensively during the measurements. Additionally, this allows for the detection of slowly diffusing entities, which is a common feature of proteins expressed in cell membranes. The analysis procedure to extract molecular concentration and diffusion parameters from the generated autocorrelation curves is highlighted. In summary, a basic protocol for FCS measurements is provided; it can be followed by immunologists with an understanding of confocal microscopy but with no other previous experience of techniques for measuring dynamic parameters, such as molecular diffusion rates.

A method to measure protein diffusion in membranes of primary immune cells using fluorescence correlation spectroscopy (FCS) is described. In this paper, the use of antibodies for fluorescent labeling is illustrated.

Fluorescence correlation spectroscopy (FCS) is a powerful technique for studying the diffusion of molecules within biological membranes with high spatial ...

Generation of Large Numbers of Myeloid Progenitors and Dendritic Cell Precursors from Murine Bone Marrow Using a Novel Cell Sorting Strategy

Cultures of monocyte-derived dendritic cells (moDC) generated from mouse bone marrow using Granulocyte-Macrophage Colony Stimulating Factor (GM-CSF) have recently been recognized to be more heterogeneous than previously appreciated. These cultures routinely contain moDC as well monocyte-derived macrophages (moMac), and even some less developed cells such as monocytes. The goal of this protocol is to provide a consistent method for identification and separation of the many cell types present in these cultures as they develop, so that their specific functions may be further investigated. The sorting strategy presented here separates cells first into four populations based on expression of Ly6C and CD115, both of which are expressed transiently by cells as they develop in GM-CSF-driven culture. These four populations include Common myeloid progenitors or CMP (Ly6C-, CD115-), granulocyte/macrophage progenitors or GMP (Ly6C+, CD115-), monocytes (Ly6C+, CD115+), and monocyte-derived macrophages or moMac (Ly6C-, CD115+). CD11c is also added to the sorting strategy to distinguish two populations within the Ly6C-, CD115- population: CMP (CD11c-) and moDC (CD11c+). Finally, two populations may be further distinguished within the Ly6C-, CD115+ population based on the level of MHC class II expression. MoMacs express lower levels of MHC class II, while a monocyte-derived DC precursor (moDP) expresses higher MHC class II. This method allows for the reliable isolation of several developmentally distinct populations in numbers sufficient for a variety of functional and developmental analyses. We highlight one such functional readout, the differential responses of these cell types to stimulation with Pathogen-Associated Molecular Patterns (PAMPs).

Here we provide a method for identifying and isolating large numbers of GM-CSF driven myeloid cells using high speed cell sorting. Five distinct populations (Common myeloid progenitors, granulocyte/macrophage progenitors, monocytes, monocyte-derived macrophages, and monocyte-derived DCs) can be identified based on Ly6C and CD115 expression.

Cultures of monocyte-derived dendritic cells (moDC) generated from mouse bone marrow using Granulocyte-Macrophage Colony Stimulating Factor (GM-CSF) have ...

In Vitro Canine Neutrophil Extracellular Trap Formation: Dynamic and Quantitative Analysis by Fluorescence Microscopy

In response to invading pathogens, neutrophils release neutrophil extracellular traps (NETs), which are extracellular networks of DNA decorated with histones and antimicrobial proteins. Excessive NET formation (NETosis) and citH3 release during sepsis is associated with multiple organ dysfunction and mortality in mice and humans but its implications in dogs are unknown. Herein, we describe a technique to isolate canine neutrophils from whole blood for observation and quantification of NETosis. Leukocyte-rich plasma, generated by dextran sedimentation, is separated by commercially available density gradient separation media and granulocytes collected for cell count and viability testing. To observe real-time NETosis in live neutrophils, cell permeant and cell impermeant fluorescent nucleic acid stains are added to neutrophils activated either by lipopolysaccharide (LPS) or phorbol 12-myristate 13-acetate (PMA). Changes in nuclear morphology and NET formation are observed over time by fluorescence microscopy. In vitro NETosis is further characterized by co-colocalization of cell-free DNA (cfDNA), myeloperoxidase (MPO) and citrullinated histone H3 (citH3) using a modified double-immunolabelling protocol. To objectively quantify NET formation and citH3 expression using fluorescence microscopy, NETs and citH3-positive cells are quantified in a blinded manner using available software. This technique is a specific assay to evaluate the in vitro capacity of canine neutrophils to undergo NETosis.

In Vitro Canine Neutrophil Extracellular Trap Formation: Dynamic and Quantitative Analysis by Fluorescence Microscopy

We describe methods to isolate canine neutrophils from whole blood and visualize NET formation in live neutrophils using fluorescence microscopy. Also described are protocols to quantify NET formation and citrullinated histone H3 (citH3) expression using immunofluorescence microscopy.

In response to invading pathogens, neutrophils release neutrophil extracellular traps (NETs), which are extracellular networks of DNA decorated with ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Image Processing Protocol for the Analysis of the Diffusion and Cluster Size of Membrane Receptors by Fluorescence Microscopy

Particle tracking on a video sequence and the posterior analysis of their trajectories is nowadays a common operation in many biological studies. Using the analysis of cell membrane receptor clusters as a model, we present a detailed protocol for this image analysis task using Fiji (ImageJ) and Matlab routines to: 1) define regions of interest and design masks adapted to these regions; 2) track the particles in fluorescence microscopy videos; 3) analyze the diffusion and intensity characteristics of selected tracks. The quantitative analysis of the diffusion coefficients, types of motion, and cluster size obtained by fluorescence microscopy and image processing provides a valuable tool to objectively determine particle dynamics and the consequences of modifying environmental conditions. In this article we present detailed protocols for the analysis of these features. The method described here not only allows single-molecule tracking detection, but also automates the estimation of lateral diffusion parameters at the cell membrane, classifies the type of trajectory and allows complete analysis thus overcoming the difficulties in quantifying spot size over its entire trajectory at the cell membrane.

Here, we present a protocol for single particle tracking image analysis that allows quantitative evaluation of diffusion coefficients, types of motion and cluster sizes of single particles detected by fluorescence microscopy.

Particle tracking on a video sequence and the posterior analysis of their trajectories is nowadays a common operation in many biological studies. Using ...

Gene Knock-in by CRISPR/Cas9 and Cell Sorting in Macrophage and T Cell Lines

Functional genomics studies of the immune system require genetic manipulations that involve both deletion of target genes and addition of elements to proteins of interest. Identification of gene functions in cell line models is important for gene discovery and exploration of cell-intrinsic mechanisms. However, genetic manipulations of immune cells such as T cells and macrophage cell lines using CRISPR/Cas9-mediated knock-in are difficult because of the low transfection efficiency of these cells, especially in a quiescent state. To modify genes in immune cells, drug-resistance selection and viral vectors are typically used to enrich for cells expressing the CRIPSR/Cas9 system, which inevitably results in undesirable intervention of the cells. In a previous study, we designed dual fluorescent reporters coupled to CRISPR/Cas9 that were transiently expressed after electroporation. This technical solution leads to rapid gene deletion in immune cells; however, gene knock-in in immune cells such as T cells and macrophages without the use of drug-resistance selection or viral vectors is even more challenging. In this article, we show that by using cell sorting to aid selection of cells transiently expressing CRISPR/Cas9 constructs targeting the Rosa26 locus in combination with a donor plasmid, gene knock-in can be achieved in both T cells and macrophages without drug-resistance enrichment. As an example, we show how to express human ACE2, a receptor of SARS-Cov-2, which is responsible for the current Covid-19 pandemic, in RAW264.7 macrophages by performing knock-in experiments. Such gene knock-in cells can be widely used for mechanistic studies.

This protocol uses fluorescent reporters and cell sorting to simplify knock-in experiments in macrophage and T cell lines. Two plasmids are used for these simplified knock-in experiments, namely a CRISPR/Cas9- and DsRed2-expressing plasmid and a homologous recombination donor plasmid expressing EBFP2, which is permanently integrated at the Rosa26 locus in immune cells.

Functional genomics studies of the immune system require genetic manipulations that involve both deletion of target genes and addition of elements to ...