Flow cytometry: principles and practices

Flow cytometry established itself as a valuable research tool several decades ago. It now has become a major asset in the diagnosis, prognosis, and monitoring of many medical disorders in addition to some commercial applications. Flow cytometers were initially large and somewhat cumbersome instruments, but have evolved into table-top, user-friendly, multi-functional ones. Over the years, clinical laboratory scientists have found additional ways to utilize flow cytometry that go well beyond simple particle counting. Today’s flow cytometers are used in cell differentiation, chromosome analysis, cellular immunology, clinical hematology, cancer diagnosis, pharmacology, and toxicology, to name a few applications. Though primarily used in monitoring immunocompromised (HIV+) patients and in the diagnosis and prognosis of different blood cancers, their utility has extended into other areas of science and industry. Understanding the science behind flow cytometry helps us to appreciate its usefulness and importance in the clinical laboratory.

Introduction to flow cytometry

The introduction of flow cytometry as a diagnostic tool has been a lengthy process.^1-4 In the nineteenth century, scientists were just scratching the surface of identifying various eukaryotic cells, but through advancements in microscopic techniques and the development of special stains and dyes, flow cytometry has become sophisticated science. (Table 1) outlines some key events that led to the development of the flow cytometer and cell sorters that are currently in use today.)

Beginning in the 1940s, a number of technological advances in flow cytometry were applied to clinical practices in order to better understand the morphology, kinetics, and structure(s) of various cell types, bacteria, and other microscopic particles. Clinical laboratory application of flow cytometry became particularly relevant with the advent of monoclonal antibodies (Moabs) as reported by Kohler and Milstein.⁵ With the addition of powerful lasers, highly sensitive photomultiplier tubes (PMTs), and computer technology, the sensitivity and accuracy of crude flow cytometers were greatly improved. Use of Moabs conjugated with a fluorescent dyes made for a very powerful diagnostic tool.⁶

Principles of flow cytometry

A basic concept of flow cytometry is the utilization of the principle of the laminar flow sheath. This occurs when a monodispersed suspension of particles (cells) is slowly injected under pressure into a faster-moving stream of fluid, the latter traveling at approximately 30 meters/second. This forms a “sheath” surrounding and aligning the cells and is the basis of the fluidic system utilizing hydrodynamic focusing.⁷ The cells flow through an orifice (100 µ for blood cells) in an almost identical trajectory, passing through a focal spot of intense illumination (laser) where optical and/or electrical signals reflect certain biologic properties of the cells (size, volume, internal characteristics). The sensing region is comprised of a quartz flow cell (chamber, cuvette) and analyzed at between 1,000 and 10,000 cells within a few seconds.²

The laser (light amplification by stimulated emission of radiation) provides an excellent intense light source for a flow cytometer. The laser light passes through several lenses, creating a tightly focused beam that passes through the sensing region (interrogation point) of a flow cell. The laser light hits the cells, causing a disruption of the laser light, and creating light to scatter in all directions. (Figure 1) This ensures that each cell passing through the focal point receives uniform illumination to produce maximum light scatter.

Light scatter is measured as both forward angle light scatter (FALS; 0.5°–2°) and as right angle light scatter (side scatter; 90^o; orthogonal). Forward angle scatter is proportional to the refractive index difference between the particle and the cell suspension media and, to some degree, particle size.² FALS travels straight through the flow cell until the laser light hits a barrier bar, blocking direct illumination that would otherwise interfere with measurements of FALS. The remainder of the FALS is measured by a photodiode or a photomultiplier tube (PMT). A PMT is more sensitive for analysis, especially if very small particles (such as bacteria) are measured.^2,7,8

A second lens is placed 90° (side scatter) to the laser light, collecting additional light scatter and fluorescence. Side scatter reflects the internal structures of the cells (morphology, N/C ratio, granularity), thus aiding in identifying various cell types. In addition, cells that have been exposed to fluorochrome-labeled, monoclonal antibodies can also be analyzed for positivity.  

Light scatter is collected from all angles and collimates it into parallel rays that hit unique dichroic mirrors (beam splitters). These mirrors are interference filters that allow certain wavelengths (unique to each fluorochrome used) to pass through while reflecting others. Each dichroic mirror has a corresponding bandpass filter unique to a predetermined wavelength which minimizes wavelength overlap. The light intensity is then measured by the PMT. All these data (FALS, SS, fluorescence) are collected and processed with the aid of computer-generated algorithms, collating the information into dot plots and histograms. These graphics reflect specific properties of the cells based on internal cell structures, size, and fluorescence or other special dye. One can then focus on specific regions of the histogram and restrict (gating) data presentation to just those parameters of interest. For example, lymphocytes, granulocytes, and monocytes can be easily segregated based on size and internal cell structures. 

In a similar fashion, smaller particles (platelets and cell fragments) can be isolated and eliminated from the data analysis by setting an observational threshold just above this point of collection.⁸ In addition, gates can be set to eliminate extraneous materials. For instance, dead cells and cellular debris can be electronically excluded, thus reducing background noise. Certain subpopulations of cells may also be gated out, allowing the user to analyze a specific subset of cells and focus on key characteristics of interest. 

In the early years of flow cytometry, only one fluorochrome (FITC) was used to label Moabs. Today, many fluorochromes are available that emit at different wavelengths. (Table 2), shows a few fluorochromes with their excitation and emitting wavelengths. This allows a user to create panels, where each Moab is conjugated with a different fluorochrome, thus providing an array of positive and negative results based on the characteristics of each monoclonal used. Multi-color flow cytometry is the current state-of-the-art diagnostic tool in flow cytometry and offers significantly more information related to subsets within an isolated population. From a single tube containing multiple antibodies, each labeled with a different fluorochrome, more data are obtained, in addition to saving technical and analytical time, reducing sample requirements, and decreasing reagent use. Many laboratories use four-, six-, eight-,10-, and even 12-color analyses to capture very specific cell populations or other determinants, generating significant amounts of useful information.^9-11

Multi-colored analyses can be extremely useful in clinical laboratory settings. However, the process is more complicated than performing single-fluorochrome analyses. In these instances compensation computer generated algorithms must be employed. Compensation is a process that corrects for spillover, i.e., one fluorochrome emitting into a detector region designated for another fluorochrome.¹² Fluorochromes can emit over a range of wavelengths; thus if one of those wavelengths is the same as that of a second fluorochrome, spillover occurs. Thus the need for unique barrier filters to help reduce erroneous cell identification.

Cell sorting

A flow cytometer can also be used to physically segregate unique cell populations. As cells pass through the orifice they trigger optical sensors, setting off an ultrasonic vibration (frequency between 20 and 60 kHz) on a piezoelectric crystal causing the jet stream to break up into droplets.⁸ The piezoelectric effect occurs when certain crystalline materials are distorted and subsequently polarized, creating a voltage across the crystal. When one of these crystals is deformed by vibration, compression, bending, or shear force on the surface of the material, a charge nearly proportional to the applied force is produced. Positive and negative charges are redistributed from their position within the crystalline lattice and expand along this piezoelectric axis. The end result is equal and opposite charges developed at each end of the crystal and can be transferred to the droplets containing the cells. This complex process requires precise timing of the vibration pulses in order to create the droplets. (Figure 2)

As each droplet (containing a cell) passes through an electrical field of two charged plates (one positive and one negative), cells can be separated based on computer-generated predetermined properties of those cells (size, positivity for fluorescence, stain, or other markers). Positively charged droplets containing the desired cells are deflected by the negatively charged plate and pass into a collecting chamber. In a similar fashion, negatively charged droplets containing the positively charged cells pass into another chamber. Cells can be sorted at the rate of about 5,000 per second with 98% purity. Some cell sorters can separate up to four populations, making this a very powerful technique to isolate specific cell types.

Anthony Kurec, MS, H(ASCP)DLM, is Clinical Associate Professor, Emeritus, at SUNY Upstate Medical University in Syracuse, New York, and a member of the MLO Editorial Advisory Board.

References

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