Considerations When Purchasing a Flow Cytometer

The capabilities of an optics system in a flow cytometer will directly influence how an experiment is designed and executed. Lasers provide the light to excite fluorophores conjugated to antibodies or reagents for use in the detection and analysis of various components and aspects of the cell.  Detectors are the devices that capture the light of specific wavelengths, amplify, and resolve this light to provide the information of the specific physical characteristic details the fluorophore is meant to measure. The technology behind the lasers and detectors dictates the number and type of fluorophores used in an experiment. This is why it’s important to understand the options of the many components that create the optical system in order to obtain optimal data.

The identification of a cell and its physical characteristics is dependent on the power of the laser beam and the sensitivity of the detection system. Most experiments require multiple fluorophores to tag cell surface and intracellular proteins in order to identify cell populations, cell cycle, population doublings, apoptosis, and cell viability. Good optics systems with quality components will capture the full-range of biologic and biochemical properties of a cell.

Each fluorophore has an excitation energy range and a specific light-emitting spectrum. Cells can be labeled with multiple fluorescence-labeled antibodies, fluorescent dyes, and fluorescent proteins. Lasers emit power at specific wavelengths to provide the different amounts of required excitation energy.

Some fluorophores have multiple excitation wavelengths of different efficiencies. For example, either a blue 488 or yellow 561 nm laser can excite the fluorophore PE. A blue 488 laser excites PE at a lower energy level. A yellow 561 nm laser provides more excitation to the fluorophore and results in a greater amount of emitted photons. Stronger emitted light may resolve a small, dim population of cells that might not have been seen on an instrument without a yellow 561 nm laser.

Commercially available flow cytometers may include violet (405 nm), blue (488 nm), green (532 nm), yellow (561 nm), and red (640 nm) lasers. Base instruments typically include the blue 488 nm laser. Adding more lasers provides the option to increase the number of fluorophores that may be used in one experiment. Violet, blue, yellow-green, and red lasers are sufficient to excite most fluorophores used in multi-fluorophore experiments. Some flow cytometers can be equipped with a UV laser in the 350 nm range to spread out the excitation and emission wavelengths when designing multicolor experiments.

Every cell should receive similar laser excitation energy in order to compare emitted fluorescent signals between cells in an experiment. Consistent laser beam output with correct laser alignment helps provide an optimal signal to every cell that passes through the interrogation point. Well-aligned laser beams result in data produced by biological, not instrument, variation.

In samples with weakly labeled cells, suboptimal excitation energy can lead to negative results. When laser power is not consistent, there will be more variation between collected emissions from cells resulting in greater statistical error. Shifts in laser alignment may provide less or unequal excitation energy to each cell passing through the beam.

Gaussian laser beams are found in most flow cytometers. Cells must pass through the center of the beam for optimal excitation energy.

Recently developed flat-top laser profiles cover more area to excite fluorophores. The wider apex allows cells in a sample to have an even application of energy to help produce consistent results.

Multiple lasers in one instrument are most commonly arranged in two formations. The arrangement affects both the number of fluorophores used in an experiment and also the detection of cell fluorescence.

Collinear spatial arrangement fires all lasers at the same time to focus laser light onto one area, resulting in the simultaneous excitation of many different fluorescent dyes.

This laser arrangement is less flexible in multicolor panel building as multiple fluorescence signals are simultaneously emitted and require greater compensation. A collinear laser arrangement works well with lasers of two different wavelengths that excite fluorescent dyes with minimally or non-overlapping emission spectra. Less expensive flow cytometer instruments may have this type of laser arrangement to save on component cost.

Spatially separated arrangement sequentially fires lasers on the cell stream. Cells stained with multiple fluorophores pass through each laser beam one at a time for separate excitation and detection of emission spectra. This type of arrangement provides flexibility for the number of dyes used in one experiment. The individually emitted fluorescence signals provides higher sensitivity and requires less compensation. Large, multicolor panels can be more readily built using this arrangement as it generates significantly less fluorescence overlap in similar emission spectra.

Excited fluorophores emit light in a limited spectrum, not at a specific wavelength. Emitted fluorescence needs to be passed through optical filters before it is captured and analyzed. Without fluorescence separation, the unfiltered light from multiple fluorophores would overlap and be difficult to analyze

Cells labeled with multiple fluorophore-conjugated antibodies can be analyzed because of the development of optical filters. These light separation components are the most commonly used for fluorescence separation in flow cytometers as they are easily accessible and user interchangeable. Combining different filters allows for varying color combinations to create both small and large multicolor panels. This is a tried-and-true system with durable components. Look for a system with easily changeable filters.

Spectral flow cytometry is different from conventional flow cytometry. Instruments with this technology use prisms to focus emitted fluorescence in the visible light spectrum. Light emitted from multiple fluorophores will overlap and requires a second step with complex algorithms for separation. The benefit of this technology is that it expands the total number of color channels from fewer lasers. This technology is still developing for flow cytometry, and computational fluorescence separation maybe limited for certain fluorophores.

The emitted fluorescence from a single cell is a weak signal. The light needs to be converted into an adjustable form of data for analysis. An emission detection device captures fluorescence light and converts the light waves into an electronic signal. This step is important as individual electronic signals can be amplified and the data can then be processed by the user.

Photomultiplier tubes (PMTs) are detection devices found in the majority of flow cytometers. Multiple PMTs are arranged at the end of the optical path and capture each filtered fluorescence signal. This type of detection device is useful with multicolor panel experiments because PMTs can capture a dynamic range of fluorescence, from very dim to very bright signals in a linear manner from the broad range of the spectrum from UV to IR.

Alternative detection devices include the avalanche photodiode (APD) to conduct and multiply electrons generated from emitted fluorescence. Experiments that heavily use fluorophores emitting in the far-red spectra can benefit from APDs. These detectors efficiently capture signals emitted in the red and near- infrared spectral regions.

Sample background may be detected as autofluorescence from cells or other components of a sample running on a flow cytometer. Fully visualizing the physical characteristics of stained cells requires capturing the full range of negative (dim) signal through positive signal. In other words, the ideal PMT voltage must be adjusted to view the maximum separation between the associated stained fluorophore and the cell’s detectable (negative) autofluorescence.

Before running cell samples, PMT voltages of the flow cytometer should be optimized using a fluorescent standard to calculate the optimal voltages for each PMT.

A PMT voltage may be adjusted if necessary, especially if the positive signal is too bright. Users may adjust the voltage by using unstained control cells as a negative for background signal. Lowering the voltage of the PMT allows proper measurement of the positive signal and decreases the intensity value of the negative population.

The amount of signal from fluorescence light detected by the flow cytometer is controlled by voltage settings. Setting a higher or lower voltage produces more or less gain on the PMT detector for proper fluorescence signal measurement.

Locked voltage settings are part of simple flow cytometer function, thus simple flow cytometers offer a limited range of detectable emitted fluorescence. Adjusting voltage settings can be challenging since the user needs to be able to distinguish positive cell populations from background. New users may over- or under-adjust voltages. To help these users, voltages are locked for a small panel of fluorescent dyes and their associated detectors. Instruments with locked settings simplify the process by limiting the ability to change voltage settings.

Unlocked or adjustable voltage settings are preferable for users running multicolor panel experiments. Locked voltage settings are preset to detect fluorescence emitted from a limited panel of fluorescent dyes. Adjustable voltage settings are not limited to a set panel of fluorescent dyes. This feature allows users to employ larger panels or include more types of dyes. Adjustable voltage settings also allow users to check their positive, single-stained controls to place their PMT signals on scale. PMT voltages may also be adjusted for optimal signal compensation as part of multiparameter flow cytometry experiments.

Emission from multiple fluorophores produces a range of light in the visible spectrum. Data collection from multicolor panels will produce overlapping fluorescence signals. This can be problematic as positive single-labeled cells may appear as double-positive cells.

Compensation is the separation of signal from each fluorophore by a set of algorithms. Traditional compensation separates specific wavelengths. The process measures each fluorophore individually and then uses algorithms to create a matrix of correction values. These corrections are applied to the spectral overlap to produce individual values. This process straightforwardly and quickly computes signal compensation.

Spectral unmixing is different from traditional compensation in that it provides the spectrum of emitting light versus a single wavelength.

Components to capture emitted light in both PMTs and APDs are called channels. The number of fluorophores detected by a flow cytometer will be determined by the number of available channels.

A one-laser system can have multiple channels.  This means a single laser can excite multiple fluorophores, and the detectors can capture emitted light at different wavelengths from a sample. This type of experiment requires complex compensation as all fluorophores can emit light in overlapping wavelengths.

Multiple lasers may be of benefit since they provide more channels. Additional channels confer easier panel design as there are more fluorophore choices and less compensation. This may allow for an experiment with better resolution of populations.