Flow cytometer is a powerful tool used for the interrogation of characteristics and phenotyping the cells. Fluorescence of cells is associated with dyes used for labeling process and it is based on the property of light scattering by cells. Flow cytometry aids in the identification of different cell types of a heterogeneous mixture and cell sorting is a major application of flow cytometer.
History of flow cytometry
In the year 1954, Wallace COulter first proposed to devise an instrument, which can electronically measure cells in a conductive liquid. This form the basis of a flow cytometer. The first one to be developed was a single cell system. In 1965, a two parameter flow cytometer that absorbed absorption and back scattering of unstained cells was developed by Kamentsy. This instrument was used to determine cell nucleic acid content and size and it was the first multi-parameter flow cytometer. The same year Fulwyler invented the first cell sorter. The Fulwyler instrument used an electrostatic deflection ink-jet recording technique that enabled the instrument to sort cells in volume at a rate of 1000 cells/s. Thompson used electrostatic charging of droplets which enhanced the development of cell sorters. Van Dilla et al. exploited the real volume differences of cells to prepare suspensions of highly purified (>95%) human granulocytes and lymphocytes. The first clinical flow cytometers were introduced in the year 1983. Following the increase in the use of computers, the rate and number of parameters increased dramatically.
Principles of Flow Cytometry
All forms of cytometry depend on the basic laws of physics, such as of fluidics, optics, and electronics. Flow cytometry is a system for sensing cells or particles as they move in a liquid stream through a laser/ light beam past a sensing area. The relative light-scattering and color-discriminated fluorescence of the microscopic particles are measured and analysed. Differentiation of the cells are based on size, granularity, and whether the cell is carrying fluorescent molecules in the form of either antibodies or dyes. As the cell passes through the laser beam, light is scattered in all directions. The light scattered in the forward direction is at low angles (0.5–10°) from the axis. This scattering is proportional to the square of the radius of a sphere and so as to the size of the cell or particle. Light scattered at may be considered to have entered the cell and be reflected and refracted by the nucleus and other contents of the cell and it represents granularity of the cell.
The cells may be labeled with fluorochrome-linked antibodies or stained with a fluorescent membrane, cytoplasmic, or nuclear dyes. Thus, differentiation of cell types, the presence of membrane receptors and antigens, membrane potential, pH, enzyme activity, and DNA content may be identified.
Flow cytometers are multiparametric. They can record several measurements of each cell. This helps in identification of a homogeneous subpopulation within a heterogeneous population. This is one of the most useful features of flow cytometers and makes them preferable to other instruments such as spectrofluorometers, in which measurements are based on analysis of the entire population. With three lasers it is possible to analyze up to 11 parameters. A typical flow cytometer consists of three functional units: (1) one or more laser light sources and a sensing system that comprises the sample/flow chamber and optical assembly, (2) a hydraulic system that controls the passage of cells through the sensing system, and (3) a computer system that collects data and performs analytical routines on the electrical signals relayed from the sensing system.
The flow chamber is instrumental in delivering the cells in suspension to the specific point that is intersected by the illuminating beam and the plane of focus of the optical assembly. Flow chambers comprise flat-sided cuvets to minimize unwanted light reflections, and, where cell sorting is required, so-called stream or “jet in air” flow cells are used. Cells suspended in isotonic fluid are transported through the sensing system. Most instruments use a lamina/sheath flow technique to confine cells to the center of the flow stream; this also reduces blockage due to clumping. Cells enter the chamber under pressure through a small aperture that is surrounded by sheath fluid. The sheath fluid in the sample chamber creates a hydrodynamic focusing effect and draws the sample fluid into a stream. An accurate and precise positioning of the sample fluid within the sheath fluid is critical, and adjustment of the relative sheath and sample pressures ensures that cells pass one by one through the detection point. Water-cooled laser sources with an output power in the range of 50 mW– 5 W may be used for fluorescence and light-scatter measurements. Air-cooled lasers have a maximum output of 100 mW. Lasers have the advantage of producing an intense beam of monochromatic light which in some systems may be tuned to several different wavelengths. The most common lasers used in flow cytometry are argon lasers, which produce light between wavelengths of 351 and 528 nm. Other lasers used, include UV lasers, which produce light between 325 and 363 nm; krypton lasers, which produce light between 350 and 799 nm; helium–neon lasers, which produce light at 543, 594, 611, and 633 nm; and helium–cadmium lasers, which produce light at 325 and 441 nm.
Fluorescence is excited as cells traverse the laser excitation beam, and this fluorescence is collected by optics at right angles to the incident beam. A barrier filter blocks laser excitation illumination, while dichroic mirrors and appropriate filters are used to select the required wavelengths of fluorescence for measurement. The photons of light falling upon the detectors are converted by photomultiplier tubes (PMTs) to an electrical impulse, and this signal is processed by an analog-to-digital (A-to-D) converter that changes the electrical pulse to a numerical signal. The quantity and intensity of the fluorescence are recorded by the computer system.
Light-Scatter and Fluorescence Detection
Light scattered by particles as they pass through a laser or light source must be efficiently detected, and fluorescent light of a given wavelength requires specific identification. Photodiodes are used as forwarding angle light (FAL) sensors; they may be used with neutral density filters that proportionally reduce the amount of light received by the detector. A beam absorber (or diffuser or obscuration bar) is placed across the front of the detector to stop the laser beam itself and any diffracted light from entering the detector. The scattered light is focused by a collecting lens onto the photodiode(s) which converts the photons into voltage pulses proportional to the amount of light collected (integrated pulse).
Fluorescence detectors are usually placed at right angles to the laser beam and sample stream. To detect the components of the beam, filters and dichroic mirrors are used to remove unwanted wavelengths of light and direct light to the correct detector(s).
Filters are used in sets, usually in pairs of a band-pass filter with a dichroic mirror or beam splitter. Beam splitters are metallic-coated quartz substrates and are designed to work at a 458 angle of incidence. Filters have numbers that indicate reflection/transmission value for the center wavelength and bandwidth (nm).
Light-scatter signals may be a measure of a combination of parameters: (1) the size (projected surface area) of the particle, (2) the surface topography rough or smooth), (3) the OD (OD will be influenced by the light absorbed and the refractive index will determine the light refracted through the particle), and (4) the internal structure of the particle (granular or uniform). The purpose of analyzing the light-scatter or fluorescence signal is to determine the difference between particles in terms of voltage output from detectors. A measure of the maximum voltage (or peak) level reached as the particle passes through the laser beam may be measured. The highest voltage level reached by the pulse (pulse height) may be a measure of the maximum fluorescence given off by a particle. Particles with different amounts of associated fluorescence have different pulse heights and therefore different peak pulses. A particle with fluorescent molecules spread uniformly over the surface will produce a wide peak pulse compared with a particle with fluorescence concentrated at one point. The latter will produce a narrower and sharper peak pulse. However, the peak of the pulse may be the same for both particles and so they become indistinguishable on the basis of this parameter. The area under the two pulses will, however, be different. The area under the pulse allows the generation of a second parameter, referred to as the integrated pulse. A third parameter may also be used if the ratio of the peak or integrated pulse is measured. This is termed time of flight.
Apoptosis is a form of cell death that has evolved to counter cell proliferation. Examples of apoptosis are seen during fetal development (e.g., loss of webbing between digits), in the deletion of autoreactive immune cells in the neonate and during the maintenance of tissue and organ homeostasis in the adult. Aberrations in apoptosis control result in disease; cancer and autoimmune diseases can originate from the inhibition of apoptosis; and degenerative diseases (e.g., Parkinson’s disease) can be a consequence of unwanted activation of apoptosis. Apoptosis contrasts with necrosis, often referred to as “accidental cell death,” in that the plasma cell membrane remains intact.
Detection of Apoptosis Using Uptake of 7-Aminoactinomycin D (7-AAD)
- Wash cells in PBS and resuspend at 1 × 106/mL in PBS.
- Add 7-AAD to 1 mL of cells (i.e., 106 cells) to give a final 7-AAD concentration of 20 μg/mL and Incubate the cells at 4°C for 20 min.
- Pellet the cells and resuspend in 500 μL of 2% paraformaldehyde.
- Analyze the cells within 30 min of fixation with a dot plot using forward scatter on the x-axis and 7-AAD fluorescence on the y-axis.
- Three distinct populations should be observed: a population with 7-AAD fluorescence equivalent to the negative control (non-stained paraformaldehyde fixed cells, which are the viable cells), 7-AAD-bright cells that are the dead or late apoptotic cells, and 7-AAD-dim cells that are the apoptotic population.
Detection of Apoptosis and Necrosis Using fluorescent dye and Potassium Iodide (PI)
- Harvest 1 × 106 cells and resuspend in 1 mL of PBS.
- Add Hoechst 33342 to a final concentration of 1 μg/mL.
- Add PI to a final concentration of 5 μg/mL.
- Incubate for 5 min and analyze using a dot blot of PI on the x-axis and Hoechst on the y-axis.
Cell Sorting by Flow Cytometry
Flow sorters have become a widespread and vital resource in the biological sciences and beyond. Their main purpose is to retrieve populations of interest from a heterogeneous population for further study. If a cell or particle can be specifically identified by its physical or chemical characteristics, it can be separated using a flow sorter. This chapter discusses the ways in which this may be done, the principles behind particle sorting, and the practicalities of a successful sorting experiment
Principles of Particle Sorting
Most analytical flow cytometers are enclosed, in that the cells are aspirated from a reservoir and hydrodynamically focused so that they pass one by one through a light source, generally from one or more lasers. At this point, scattered light and fluorescence signals are generated, detected, and measured. After this, cells are removed under vacuum to a waste reservoir. In general, flow sorters use a principle involving the electrostatic deflection of charged droplets similar to that used in inkjet printers. To sort particles by this method, the process has to be performed in a more open system where cells are ejected into air in a stream of sheath fluid.
Any fluid stream ejected into air will break up into droplets but this is not a stable process; the distance from the orifice that the stream begins to break up will depend on many factors such as the orifice size, the pressure of the sheath fluid, the ambient temperature, and the viscosity of the fluid. However, if a stationary wave of vibration of known frequency and amplitude is applied to the fluid stream, it is possible to stabilize the break-off, and for a given set of conditions, the size of the droplets and the distance between drops will also stabilize. In a flow sorter, this vibration is produced by a transducer, which is generally a piezoelectric crystal acoustically coupled to the nozzle. As cells are ejected from a nozzle, they pass through one or more laser beams and at this point – the moment of analysis – information is gathered about the cell or particle.
The distance, and therefore time, between the point of analysis and the point at which the cell breaks off from the solid stream in a droplet, is constant and under given conditions can be calculated. This distance between the laser intercept and the break-off point is measured in drop equivalents and is often referred to as the drop delay. The calculation and monitoring of this are critical and is the factor that makes a sort successful or not. The drop break-off can be observed microscopically under stroboscopic illumination to allow the break-off point to be monitored. The drop delay is calculated by determining how many drops are in the distance between the analysis point and the break-off point; drops will start to form as soon as the stream emerges from the nozzle but will be coalesced until the break-off point. There are several ways of measuring the drop delay: by counting the number of drops formed at a known distance, by sorting beads onto slides at varying drop delays and checking microscopically, or by viewing fluorescent beads in sorted side streams. The precise way of calculating drop delay will vary with the type of sorter used. Once the drop delay is calculated, it is possible to charge through the stream at the precise moment that the first drop is forming. Therefore, individual drops, as they break away from the solid stream, can be independently charged and will carry a positive charge, a negative charge, or will remain uncharged. The individual drops then pass through a static electrical field created by two charged plates. The voltage between the plates will be in the range of 2000 to 6000 V depending on the flow sorter used and the number of populations required from the sort. Charged drops are attracted to the plate of opposite polarity and will be deflected into collection vessels. The formation of drops and the determination of the drop delay are the factors that enable the flow sorter to be able to sort cells.
The speed of sorting depends on the time taken to generate a droplet, which depends on the frequency of the transducer. The frequency of droplet production in a stream-in-air sorter is determined by the jet velocity and the jet diameter and is defined as f = v/4.5 d, where v is the velocity of the fluid and d the diameter of the orifice.
Sample preparation prior to sorting is important; Successful sorting depends almost entirely on the state of the input sample. It is a prerequisite for flow cytometry that cells or particles be in a monodispersed suspension. This is relatively easy when the cells used are in a natural suspension (e.g., blood cells or suspension cultured cells) but more problematic when using cells from adherent cultures or cells from solid tissue. However, there are several well-established methods for preparing samples for flow sorting.
Preparation from Suspension Cells
- Take cells directly from a culture flask into 50-mL conical tubes and centrifuge at 400g for 5 min.
- Discard the supernatant and re-suspend in medium (cell culture medium or phosphate– buffered saline [PBS] with 1% bovine serum albumin).
- Centrifuge again at 400g and discard supernatant. Count cells and re-suspend at an appropriate concentration, which will vary with sorter used but will be in the range of 1 × 106 – 1 × 107 per mL.
Preparation from Adherent Cells
- Harvest cells by using trypsin (0.25% w/v) or Versene (0.2% w/v). Transfer cells to 50-mL conical tubes and centrifuge at 400g for 5 min.
- Discard the supernatant and re-suspend in medium (cell culture medium or PBS with 1% bovine serum albumin).
- Centrifuge again at 400g and discard supernatant. Re-suspend the cells in a small volume of medium and aspirate up and down through a pipette several times to help disaggregate clumps. Count cells and re-suspend at an appropriate concentration.
Preparation from Solid Tissue
- Place tissue in a sterile Petri dish. Tease tissue apart using a needle and scalpel or alternatively use an automated system. In addition, enzymatic disaggregation (e.g., collagenase (220 U/mL) may also help free single cells.
- Decant cells into a 50-mL conical tube and centrifuge at 400g for 5 min.
- Discard the supernatant and re-suspend in medium (cell culture medium or PBS with 1% bovine serum albumin).
- Centrifuge again at 400g and discard supernatant. Re-suspend the cells in a small volume of medium and count cells as above.
All preparations may be filtered through sterile nylon mesh prior to sorting; a range of pore sizes from 20 to 70 μm will be suitable for most cell types encountered.
Flow Sorter Setup
Run 70% ethanol through all fluidic lines for 30–60 min before flushing this with distilled water (30 min) and finally sterile sheath fluid (at least 30 min before commencing a sort). At all times, fluids pass through a 0.22-μm filter immediately after leaving the sheath tank. The next consideration is nozzle size. The cell type will influence the size of orifice used. A general rule of thumb is that for blockage-free sorting and coherent side streams a cell should be no more than one fifth the diameter of the nozzle.
Once aligned, the frequency and amplitude of the drop drive are altered to achieve a minimum stable break-off with coherent side streams using a test-mode sort. Although a given nozzle size has an approximate resonance frequency, there will be a variability between individual nozzles of a given size and there will also be variability on a day-to-day basis depending on conditions.
The setup of a sort depends entirely on which cells or particles are needed. As already noted, it is possible to sort according to antigen expression, fluorescent protein expression, nucleic acid content, or functionality. Although hydrodynamic focusing will help align particles in the center of a stream and to some extent keep them separate, some particle doublets will always be present. This can have a deleterious effect on sorting purity. The population or populations to be sorted are identified initially on the basis of their fluorescence characteristics. Dead cells are excluded on their positivity for the viability dye, the doublets are excluded as far as possible according to their pulse width, and finally, debris and events, which are clearly not intact cells, are excluded on the basis of their forward and 90° scatter signals. As a rule of thumb, dot plots should be used to set up sort parameters. Once the population to be sorted has been identified, the sort mode to be used has to be decided. In practice, there is a balance between purity, recovery, and yield of the sort. In the majority of sorts, only one drop per event will be sorted; this will give the greatest purity because drops containing unwanted cells would not be sorted, which is the usual requirement in a sorting experiment.