Post A – Instrument and sample preparation
When setting up a flow cytometry experiment, there are some key elements that you need to take into consideration upfront. Firstly, you need to know your instrument, optical setup, and choice of fluorochromes that can be used on your instrument. Secondly, you need to think about your test sample – what type of sample are you working with, what do you want to detect on these cells, are the markers of interest extra- or intra-cellular? In this post, we will elucidate some of the nuances of flow cytometry and provide guidance on instrument setup and sample preparation.
Flow cytometers are becoming increasingly sophisticated devices that combine fluidics, optics, electronics and software to enable detection and analysis of microscopic particles within a few seconds. The sample is aspirated into the instrument where it passes through the flow cell. Sheath fluid exerts hydrodynamic pressure on the sample, forcing it into a single cell stream, which then passes the laser sources for excitation. Light scatter (size and complexity) and fluorescence are filtered and captured by a series of detectors for conversion to electrical information which is then analysed using appropriate software. There are many different flow cytometers available for different applications, from simple bench-top 2-colour instruments to multi-laser multicolour parameter flow cytometers and cell sorters.
Laser Excitation and Emission
Most flow cytometers use more than one laser source for light excitation. Emission occurs when excitation light is absorbed and then re-radiated with no change in wavelength. Scatter is detected in the forward direction (particle size) and sideways (particle complexity). Fluorescence occurs when a molecule excited by light at one wavelength emits light of a longer wavelength, which is then blocked or filtered. Several different cellular parameters can be measured simultaneously from several different compounds being bound to one or many of the cells in the sample.
Light travels on waves that determine the colour of the light. High-energy light has shorted waves than light with less energy (Figure 1). As can be seen in Figure 1, very high-energy light such as ultraviolet (UV) light has very short wavelengths, which is why the human eye is not able to see it. Visible light has longer wavelengths (from 400 to 700nm) and produces colours of violet, blue, green, yellow, orange and red. Above 700nm is the infrared range (Figure 1). Fluorochromes are selected based on their ability to fluoresce within the range of wavelengths of light produced by the lasers. These wavelengths of lights are detected by electronic photodetectors within the flow cytometer. Fluorochromes are then conjugated to antibodies that are designed to bind to particular cellular proteins. Using more than one antibody and fluorochrome allows the researcher to perform multicolour flow cytometric analysis and characterise distinct cell populations.
An important consideration when conducting multicolour experiments is the possibility of fluorescence interference caused by dyes or fluorochromes that have overlapping emission spectra. The more fluorochromes that are being used in an experiment, the more difficult it becomes to minimise spectral overlap and prevent bleeding of fluorescence into other channels. Best practice is to spread the fluorochromes over multiple lasers in order to minimise this.
In order to minimise spectral overlap, a process known as colour compensation needs to be performed. This can be done using compensation control beads, which contain both positive and negative controls beads. These are incubated with the fluorochrome of choice. Negative controls beads will not bind the antibody whereas the positive control beads will. Both negative and positive control peaks must be easily and discreetly identified (Figure 2).
When using tandem dyes (dyes that are comprised of more than one fluorochrome to create a unique fluorescence emission), you must run compensation controls using the same fluorophore for each experiment. Examples of tandem dyes include PE-Cy5, PerCP-Cy5, and APC-Cy7.
Once single-stained compensation controls have been acquired and the data saved, compensation software can mathematically remove fluorescence overlap to simplify data interpretation and distinguish cell populations.
No matter what the end goal is for your flow cytometry experiment, it all begins with good sample preparation to ensure that you have a good single-cell suspension. Pre-analytical variables can have a huge impact on the outcomes of your experiment.
The best place to start is with a sample that is alive. This is why the time between sample collection, sample preparation and analysis needs to be as short as possible. High viability, good retention of cell surface markers, and low levels of debris are key to obtaining high-quality data.
Cells that are already dead or dying have different levels of antigen expression compared to healthy living cells. During sample preparation, the sample should always be handled gently to prevent unnecessary cell death. Harsh enzymatic treatments, vortexing and centrifugation, or prolonged lysing or fixation treatment should be avoided. The impact of leaving cells on ice and freeze-thaw cycles should also be taken into consideration. Cell debris can also result in high levels of autofluorescence as well as introducing experimental artefacts.
Validating which is the correct anticoagulant (e.g. EDTA/Heparin/SST) to collect your sample in is vital. These anticoagulants can affect the creation of a single cell suspension and binding of antibodies.
Fluctuating cell concentrations can also affect results, and can make comparisons difficult. Try to maintain a constant cell concentration in your experiment. Samples with a high cell concentration can make single-cell analysis difficult and may clog the flow cytometer, while low concentrations can lengthen acquisition time.
Prevent non-specific binding of antibodies to the cells. Some cells, like dendritic cells or monocytes, are particularly “sticky” when it comes to antibody binding. This is because these cells express a range of Fc receptors that can bind antibody, rather than their antigen-specific binding site. By incubating samples with serum from the same species as the antibodies are derived from or using blocking solutions can prevent non-specific antibody binding.
Lysing, Fixation and Permeabilization
In order to remove red cells and debris from the sample, cell lysis is performed. The best lysis procedure should be determined for each application. Various procedures that can be used include: no lyse, no wash; lyse, no wash; and lyse, wash. Over-lysing can also affect sample integrity and viability, so selection of the correct lysis buffer is key.
In some cases after lysis, a fixative may be added or cells may be permeabilized for intracellular staining. Fixation and permeabilization should be optimised like any other part of the experiment setup, and purchasing ready-to-use reagents like Instrastain (Agilent cat. no. K2311) can help give consistent results. Such reagents also come with a recommended protocol to follow, which is extremely helpful. Overfixation or over-permeabilization can affect the integrity of the sample, and can cause staining to be compromised. Insufficient permeabilization can cause insufficient intracellular staining.
- Dako – Flow Cytometry Educational Guide, 2nd Edition
- https://www.biocompare.com/Editorial-Articles/355547-How-to-Design-Successful-Flow-Cytometry-Experiments/ (accessed 9 July 2020)
- https://www.biocompare.com/Editorial-Articles/561165-Flow-Cytometry-Sample-Prep-Mistakes-to-Avoid/ (accessed 9 July 2020)
- https://www.enzolifesciences.com/science-center/technotes/2015/february/10-tips-for-successful-flow-cytometry/ (accessed 3 Aug 2020)
- https://www.aceabio.com/wp-content/uploads/Tips-and-Tricks-for-flow-cytometry.pdf (accessed 3 August 2020)