Isolation of mononuclear cells from peripheral blood and separation ...

Lab 2 Isolation of mononuclear cells from peripheral blood and separation into subpopulations

Supervisors: Sissela Broos Niclas Olsson

[email protected] [email protected]

tel: 222 96 78 tel: 222 94 13

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Aim: To gain an insight into the purification of lymphocytes and analysis of antigen expression on different cell populations.

Flowchart Filter material Ficoll-paque Lymphocytes and monocytes (PBMC) Binding to plastics Rosetting Adherent cells (monocytes) T-cells fraction B-cell fraction (incl. monocytes)

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Introduction Blood contain different types of cells, for example red blood cells (erythrocytes), which transport oxygen to all tissues in the body and white blood cells (leukocytes), which are part of the immune system. In this lab we are going to look more closely at some of the white blood cell populations. We will do so using flow cytometry which is a common technique for analysis of cells in clinical laboratories. In order to perform this analysis, we must first separate different types of leukocytes from each other by using the unique characteristics of the cells. We will isolate T-lymphocytes, monocytes and B-lymphocytes. The aim of the lab is to give an insight into different cell separation methods, as well as how to analyze cells using flow cytometry. We will use a so called ”buffy coat”, which is the remaining part when red blood cells are separated from the blood plasma in human blood. The “buffy” is obtained from the blood bank at Lund University Hospital. During day 1, we will carry out inital preparations to separate leukocytes from blood and subsequently isolate the different cell populations (monocytes, B-cells and T-cells). First, we will dispose off the red blood cells by performing a density gradient centrifugation on Ficoll-Paque. The PBMCs (peripheral blood mononuclear cells) will form a layer in the tube that will be further purified. Monocytes will be isolated by utilizing their ability to bind to plastic surfaces. Tlymphocytes will be separated using activated erythrocytes from sheep, which bind to a molecule on the surface of T-cells. The T-cell depleted fraction will then contain both B-cells and monocytes. The different cell populations will finally be studied using flow cytometry on day 2.

All donors are tested before becoming approved blood donors, but there is still a risk that the material may be contagious when we receive it, so it is vital to work very carefully. Important to also bear in mind when working with cells: 1. Always keep the cell suspensions in tubes on ice. 2. Resuspend a cell pellet in a small volume of liquid first and then add more liquid. Otherwise the cells will form “clumps”. 3. Keep the lymphocytes at the right pH (6,8-7,4), even though they endure low pH better than high pH. 4. Work gently with the cells, they are small living creatures. Do not pipette quickly.

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Isolation of mononuclear leukocytes In blood transfusions it is important that the blood does not contain any white blood cells. These are separated from the blood by filtration and the collected cells are referred to as filter material (formerly known as buffy coat, which is separated by centrifugation). From this you can obtain mononuclear cells using a density gradient centrifugation on Ficoll-Paque (density = 1.078). Differencies in density will separate lymphocytes from other blood cells. After the centrifugation three or four fractions are visible:

Centrifugation

Plasma

Blood sample Lymfoc., monocyt. & tromb. δ < 1.078 Ficoll-Paque Ficoll-Paque Granulocytes δ > 1.078 Aggregated erythrocytes δ > 1.078

Counting cells in a Bürker chamber 4

The proportion of viable cells can be determined by staining the cells with trypan blue. Viable cells will not become stained while dead cells will become blue. The cells have to be counted within 3-4 minutes or the viable cells will also take up the colour. Bürker chambers and cover slips should be properly cleaned in water and alcohol. The cover slip is attached to the glass by humidifying the borders of the glass. Dilute the cells in trypan blue and let the cell suspension slip in under the cover, into one of the chambers. Abundant cell suspension should be dried off using a Kleenex. Each field in the counting chamber is divided into 9 squares separated with triple lines and such a field (A-square) contain 0,1 µl. The A-square is further divided into 16 parts, B-squares (see figure below). When counting cells in an A-square, all cells that touch the right and the upper border should be included while cells on the left and bottom line should be excluded. Count at least a hundred cells (sometimes you may have to count more than one A-square) and use the average to determine the concentration of cells. Amount of viable cells/ml=viable cells in one A-square x 104 x dilution factor

B-square

A-square

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Flow cytometry Flow cytometry is a mean of measuring certain physical and chemical characteristics of cells or particles as they travel in suspension one by one past a sensing point. The modern flow cytometer consists of a light source, collection optics, electronics and a computer to translate signals to data. In most modern cytometers the light source of choice is a laser, which emits coherent light at a specified wavelength. Scattered and emitted fluorescent light is collected by two lenses (one set in front of the light source and one set at right angles) and by a series of optics, beam splitters and filters to allow specific bands of fluorescence to be measured. Physical characteristics such as cell size, shape, internal complexity and any cell component or function that can be detected by a fluorescent compound can be examined. So the applications of Flow Cytometry are numerous, and this has led to the widespread use of these instruments in the biological and medical fields.

The term "Flow Cytometry" derives from the measurement (meter) of single cells (cyto) as they flow past a series of detectors. The acronym FACS (Fluorescence Activated Cell Sorting) and Flow Cytometry are used interchangeably. The fundamental concept is that cells flow one at a time through a region of interrogation where multiple biophysical properties of each cell can be measured at rates of over 1000 cells per second. These biophysical properties are then correlated with biological and biochemical properties of interest. The high through-put of cells allows for rare cells, which may have inherent or inducible differences, to be easily detected and identified from the remainder of the cell population.

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In order to make the measurement of biological/biochemical properties of interest easier, the cells are usually stained with fluorescent dyes, which bind specifically to cellular constituents. The dyes are excited by the laser beam, and emit light at a longer wavelength. Photo Multiplicator Tubes (PMT) detects the emitted light as an analogue electronic signal, which subsequently is converted to digital information to be analyzed in a computer. Three types of data are generated: Forward scatter (FSC) Approximate cell size Side/orthogonal scatter (SSC) Cell complexity or granularity Fluorescence Fluorescent labeling is used to investigate cell structure and function Scatter Forward and side scatter are used for preliminary identification of cells. In a peripheral blood sample, lymphocyte, monocyte and granulocyte populations can be defined on the basis of forward and side scatter. Forward and side scatter are also used to exclude debris and dead cells. Fluorescence Labeling cells with fluorescent dyes allows investigation of structure and function. Fluorescence intensities are typically measured at several different wavelengths simultaneously for each cell. Fluorescent probes are used to report the quantities of specific components of the cells. Fluorescent antibodies are often used to report the densities of specific surface receptors, and thus to distinguish subpopulations of differentiated cell types, including cells expressing a transgene. By making them fluorescent, the binding of viruses or hormones to surface receptors can be measured. Intracellular components can also be reported by fluorescent probes, including total DNA/cell (allowing cell cycle analysis), newly synthesized DNA, specific nucleotide sequences in DNA or mRNA, filamentous actin, and any structure for which an antibody is available. Flow cytometry can also monitor rapid changes in intracellular free calcium, membrane potential, pH, or free fatty acids. Immunofluorescence, the most widely used application, involves the staining of cells with antibodies conjugated to fluorescent dyes such as fluorescein (FITC) and phycoerythrin (PE). This method is often used to label molecules on the cell surface, but antibodies can also be directed at intracellular targets in the cytoplasm. In direct staining the antibody is directly conjugated to a fluorescent dye (e.g. anti-CD4 PE). Cells are stained in one step. In indirect staining the primary antibody is not labeled. A second antibody conjugated to a fluorescent dye, and specific for the first antibody is added. For example, if the anti-CD4 antibody was a mouse IgG then the second antibody could be a rat antibody raised against mouse IgG.

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Analysis The ability of Flow Cytometers to evaluate cells at an extremely rapid rate (e.g. up to 20,000 events per second) makes this technology ideally suited for the reliable and accurate quantitative analysis of selected physical properties of cells of interest. The sensitivity of these instruments for detecting the presence of molecules expressed at low levels is impressive; given high quality cell preparations and reagents, as few as 500 molecules per cell may be detected. At the Department of Immunotechnology the Becton-Dickinson FACS Canto II will be used for analysis. The BD FACSCanto system is built with blue (488 nm, air-cooled, 20 mW solid state) and red (633 nm, 17 mW HeNe) excitation sources. The laser beams are then routed via fiber optics to the beamshaping prisms where two lasers are projected onto separate spots in the flow cell. Here, the particles intercept with the laser excitation beams and optical signals are collected at the backside of the flow cell with a gel-coupled collection lens. The system has 6 different colour detectors and several different dyes can thereby be used. In addition the system has a very fast acquisition rates (up 10,000 events per sec).

Cell sorting One of the properties of the larger flow cytometers is the ability to electronically deflect cells with preset, defined properties into a separate collection tube. In these instruments the fluidics hydrodynamically focuses the cell stream to within an uncertainty of a small fraction of a cell diameter and break the stream into uniform-sized droplets to separate individual cells. The electronics quantitate the faint flashes of scattered and fluorescent light, and, under computer control, electrically charge droplets containing cells of interest so that they can be deflected into a separate test tube or culture wells. For cell purification, flow cytometry is especially well suited for applications requiring high purity. Because multiple fluorochromes (e.g. up to five distinct fluorescent probes reacting with different cell associated molecules) can be assessed simultaneously, cell sorting by flow cytometry can separate complex mixtures of cells on the basis of multiple marker expression. At the Department of Immunotechnology we use a Becton-Dickinson FACS Aria for cell sorting. This Aria is a 9 detectors instrument with 3 lasers and digital acquisition rates of up to 70,000 events/second. The bitmapped sorting capability of the Aria digital electronics allows the operator to set up sophisticated sort decisions using logical gates combined with the Boolean operators AND, OR, and NOT. The Aria can also be used for multicolor assays with too many colors (nine) for the other instrument.

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Important! The laboratory manual is only a guide. Some changes may occur during the lab and these must be documented and described in the lab report. DAY 1: 1.

Fill a T75:a with 250 ml PBS. Cut the end of the filter tubing that is least blood filled and connect it to the vacuum tubing. Cut the other side and place it in the T75 with PBS. Turn on the vacuum and wait until all PBS has gone through the filter. Turn the vacuum off.

2.

Disconnect the bucket from the vacuum tubing and carefully put it in the LAF bench. Carefully swirl the bucket and remove the top. Transfer the blood solution into 6 x 50 ml Falcon tubes using a 50 ml stripette. Fill up the last tube with PBS to balance in the centrifuge.

3.

Centrifuge the tubes at 1800 rpm for 5 min (room temp.)

4.

Aspirate the supernatant and leave about 10 ml in each tube. Pool the solution into two of the tubes and add PBS for a total volume of 75 ml.

5.

Place 15 ml of Ficoll Paque into three 50 ml tubes/group.

6.

Carefully transfer 25 ml of the diluted filter material ONTO the Ficoll Paque so that they form two separate layers.

7.

Centrifuge in room temperature at 1800 rpm for 20 min without breaks.

8.

The leukocytes can now be collected at the interphase between the Ficoll Paque and the plasma (se figure at page 4). Transfer them carefully using a 10 ml pipette or a pasteur pipette into four new 50 ml tubes. Try to avoid aspirating the plasma and Ficoll Paque.

9.

Dilute the cell suspensions with PBS up to 50 ml in each tube.

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10.

Centrifuge 1250 rpm for 10 min (room temperature).

11.

Aspirate the supernatants and be careful not to lose the cells. Resuspend the pellet BEFORE PBS is added.

12.

Resuspend the cells in 10 ml cold PBS and pool the cells into one tube. Always keep the cells on ice after this point.

13.

Centrifuge the cells at 900 rpm for 10 min (4°C). The leukocytes will now form a pellet while the trombocytes will stay in the supernatant.

14.

Carefully aspirate the supernatant (the pellet is loose!!!). Resuspend the cells in 10 ml PBS, take out a sample and count the cells (dilute 50 times before counting) (see page 5). Repeat the washing step (number 10) while you are counting.

15.

Resuspend the cells in cell culture medium (R5) to a cell concentration of 10 x 106 cells/ml.

16.

Take out 3 times 50µl samples and transfer them to 3 different 5 ml tubes (”FACS tubes”) (=0,5 x 106 cells). Mark them with A1, A2 and A3 and the name of the group and keep them in the fridge before the analysis on day 2.

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Binding of adherent cells to plastic surfaces The cells (from step 12) that we have at this stage are lymphocytes and monocytes. Monocytes could be separated by plastic adhesion, since they adhere to plastic surfaces and lymphocytes do not. Take ~ 75 x 106 cells (