Immunology

Experiment 10

Aim of the Experiment

Demonstration of Fluorescence-activated Cell Sorting (FACS) in order to identify specific cell populations by immunofluorescent detection.

Introduction

Cytometry, is the measurement of cell characteristics, which can include cell size, cell count, cell cycle and more. Flow cytometry is a cell analysis technique that was first used in the 1950s to measure the volume of cells in a rapidly flowing fluid stream as they passed in front of a viewing aperture. This technique allows researchers to get highly specific information about individual cells. Because of its speed and ability to scrutinize at the single-cell level, flow cytometry offers the researchers the statistical power to rapidly analyze and characterize millions of cells. The most critical requirement for efficient and effective flow cytometry analysis is that the sample be a single-cell suspension. This helps ensure that every cell is analyzed independently.

An additional capability of specialized flow cytometers is the ability to sort cells and recover the subsets for post experimental use. This specialized flow cytometer is called a Fluorescence activated cell sorter (FACS), a term that is sometimes erroneously used interchangeably with ‘flow cytometer’. This usage is incorrect. A flow cytometer is an analytical machine that does not perform cell sorting. Cell sorters use fluidics and fluorescence components similar to those in flow cytometers, but are able to divert a specific population from within a heterogeneous sample into a separate tube, typically based on specified fluorescence characteristics. If collected under sterile conditions, these cells can be further cultured, manipulated, and studied.

Principle

Fluorescence-activated cell sorting (FACS) is a specialized type of flow cytometry. It provides a method for sorting a heterogeneous mixture of biological cells into two or more containers, one cell at a time, based upon the specific light scattering and fluorescent characteristics of each cell. It is a useful scientific instrument, as it provides fast, objective and quantitative recording of fluorescent signals from individual cells as well as physical separation of cells of particular interest.

how a flow cytometer works

The three main components of a flow cytometer are the fluidics, optics, and electronics.

• The fluidics system of a flow cytometer is responsible for transporting sample from the sample tube to the flow cell. Once through the flow cell (and past the laser), the sample is either sorted (in the case of cell sorters) or transported to waste.

• The components of the optical system include excitation light sources, lenses, and filters used to collect and move light around the instrument and the detection system that generates the photocurrent.

• The electronics are the brains of the flow cytometer. Here, the photocurrent from the detector is digitized and processed to be saved for subsequent analysis.

Sample uptake into the instrument

A typical experiment begins with fluorescently labeled cells in a single-cell suspension, but a sample containing particles of any kind can be used. Once the sample is placed on the flow cytometer, the sample is taken up into the instrument, and the cells are surrounded by a physiological buffer called sheath fluid. The fluidics system—the tubing, pumps, and valves—organizes the initial sample suspension into a single-file stream of cells as they make their journey through the flow cytometer for analysis.

Interrogation point

The place where the cells interact with laser light is called the interrogation point. You might also hear it referred to as the laser intercept. This is where the action takes place. When the laser light beam illuminates a single cell, some of the light will strike physical structures within the cell, causing the light to scatter. This light scatter can be measured and correlated with relative cell size and structures inside the cell. These measurements are termed forward angle scatter(FSC) and side angle scatter (SSC), depending on where the light is collected with respect to the path of the laser. Nearly simultaneously, light from the laser will excite all fluorophores associated with the cell, which produces a fluorescence emission. All of this light is collected by the detector and processed through the electronics component of the flow cytometer. After passing through the interrogation point, the cell is no longer needed and is carried by the fluidics system to the waste container. If this were a cell sorter, this would be the point where the cell would be passed to a collection tube and used for further experiments.

Materials required

• Cell lysis buffer

• 5 mL FACS tubes (Falcon)

• Block

• FACS Staining Buffer (1XPBS w/ 3% calf serum and 0.05% sodium azide)

• Sorting Buffer (1xPBS w/ 0.1% BSA or 0.5% FCS)

• Collection Buffer (depends on application, RPMI or PBS/serum)

• 70uM filter

Procedure

Preparation of cell suspensions

1. Obtain cell suspensions.

a. For non-adherent cell populations, wash cells (resuspend in buffer, centrifuge at 400 x g for 5 minutes, aspirate buffer, and resuspend in an appropriate volume of fresh buffer) in FACS staining buffer, resuspend and resuspend in a small volume of buffer.

b. For adherent cell populations, wash cells (similar to a media exchange) in FACS staining buffer and harvest cells by gently scraping the dish, plate, or culture flask. Avoid trypsin if possible as it may damage cell surface proteins. Collagenase or similar may be used if scraping is not sufficient for recovering adherent cells. Immediately wash cells (as described in 1a) again and resuspend in a small amount of FACS staining buffer.

c. For tissue samples, obtain a cell suspension homogenizing tissue in staining buffer by pressing the sample through a fine mesh sieve (nylon mesh) using a clean syringe plunger from a 3cc syringe, or similar instrument. This procedure will provide sufficient homogenization for most tissues, but other enzymatic methods are available for difficult samples.

2. Count Cells.

a. Gently mix cell suspension to ensure a homogenous mixture and reserve 20-100 ul to count cells.

b. Use a trypan blue exclusion stain or similar to exclude dead cells. Obtain cell counts using a hemocytometer or automated cell counter.

3. Resuspend cells to an appropriate concentration in FACS staining buffer. 1 x 10⁶ cells is commonly used for antibody labeling of most cell types.

Antibody Selection

1. If possible, always use directly conjugated antibodies in flow cytometry to facilitate multicolor staining and reduce background.

2. Use antibodies conjugated to bright fluorophores like PE and APC for targets that are expressed at low levels.

3. Try combining fluorophores that are on instrument channels that are far away from each other as much as you can. For instance use antibodies conjugated to fluorophores that are excited by different lasers. If combining antibodies conjugated to fluorophores on the same laser or array of your flow cytometer, try to select fluorophores with emission maxima as far as possible from one another.

Antibody Staining

Note: Fc receptors on many immune cells may bind antibodies and create false positive signals. Anti-CD16 + anti-CD32 antibodies are commonly employed as an Fc-block and may be used to reduce or eliminate this source of noise.

1. Based on the number of cells you will require, place the appropriate volume of cells suspended in FACS staining buffer into FACS tubes.

2. Add your primary antibody (preferably directly conjugated) to each sample. The dilution of antibody should be experimentally determined by running a pilot experiment on several samples. Too much antibody creates high background fluorescence and too little results in dim positive cells. An optimal antibody concentration can be found by comparing staining index at multiple concentrations and selecting the conditions that result in highest signal to noise.

3. Multiple primary antibodies may be added at the same time in this labeling step for multicolor experiments. Be sure to stain appropriate controls as well.

4. Incubate on ice for 30 min to 1 hour in the dark (ie. in a drawer in your lab).

5. Wash 1-3 times by resuspending cells in a larger volume of staining buffer (1-2ml for tubes), gently mixing, centrifuging samples at 400 x g for 5 minutes and aspirating supernatant.

6. Resuspend in an appropriate volume of staining buffer. Higher concentrations of cells may yield faster data acquisition on the flow cytometer but may also lead to formation of cell aggregates which should be avoided.

7. Proceed to running samples on the flow cytometer.

Reference Material

• FACS-Virtual Lab

• Flow cytometry

• FACS Protocol

• Flow Cytometry Animation

Questions

Q.1. What should you consider when choosing fluorophores for flow cytometry?

Q.2 How are the cells sorted?

Q.3 What is the purpose of FACS sorting?