3 Requirements For Accurate Flow Cytometry Compensation

For those new to flow cytometry, compensation is confusing at best and terrifying at worst. Likewise, those who have been doing flow cytometry since the analog ages may be holding on to practices that, while suited to the analog instruments, should be left to the annals of history. As such, a lot of time is spent discussing compensation and the best practices for this critical process.

There are 3 rules that guide proper compensation, and they’ve been written about extensively since they first appeared in the “Daily Dongle” in 2011. It is always good to review and, importantly, there are some caveats and assumptions baked into the rules which bear closer examination.

Compensation Rule 1: “Controls must be at least as bright or brighter than the sample to which the compensation will be applied.”

To ensure that the correct compensation value is calculated, we need accurate measures of our controls.

We use the slope of the line between 2 populations with different intensities in the channel of interest to calculate compensation. In theory, that calculation should yield the same result regardless of where the populations fall in the detector range. However, this is not the case in practice because there is generally greater error in the dim cell measurement than in the bright cells (Figure 1).

Figure 1: Compensation using dim or bright particles. As can be seen from this plot, if the dim particles were used for compensation at the intensities above, this value would be undercompensated. Axes are labeled with excitation line (B=488 nm) and the bandpass filter in front of the PMT.

Tacitly included in this rule is the requirement that the signal must be on-scale and in the linear range of the detector.

When the detector converts photons into photocurrent, it is important that this current is linear and proportional to the input. At voltage extremes, this relationship does not hold, so it is imperative that these bounds are determined and the signal maintained within them.

Likewise, it’s important to keep the signal on-scale. As shown below for the PE detector, the linear bounds are highlighted in yellow, and the upper scale in blue.

Figure 2: Linear bounds and upper limit of the scale for a PE labeled bead measured on a PE detector. Axes are labeled with the excitation line (G=532nm) and the bandpass filter in front of the PMT.

Thus, the first rule focuses on the expression level of the fluorochrome. Determining the best voltage for these controls is an important process in the development and validation of a panel.

Compensation Rule 2: “Background fluorescence should be the same for the positive and negative controls.”

This means that the autofluorescence of the carriers must be matched. The choice to use carrier cells or antibody capture beads depends primarily on 2 factors:

  1. Availability of cells
  2. Expression characteristics of the target antigen

If there are abundant targets on the surface of the cells and you have lots of extra cells, then using cells is no issue.

However, if the targets are not abundant (say on rare events or antigens with low levels of expression), using an antibody capture bead (ABC) is preferred.

ABCs allow you to save your cells for experimental tubes and capture large amounts of antibody to ensure the signal is at least as bright as the experimental (Rule 1), and it is the exact same fluorochrome as used on your sample (Rule 3).

To adhere to the second rule, it is important to avoid the use of the “Universal Negative” — an unstained sample (usually cells) that is collected separately from the positive control tubes and is used to set the background for compensation.

Figure 3: Comparison of using either beads or cells as the negative control for compensation. As shown in each of these plots, the beads were used for the positive control, and either beads or cells were used for the negative control. The lines help to visualize the slope of each uncompensated combination. Axes are labeled with excitation (B=488nm; R=633nm) and the bandpass filter in front of the PMT. The blue line represents the approximate slope between the negative (cells) and positive (beads), while the red line represents the approximate slope between the negative (beads) and positive (cells).

As shown in Figure 3, using the incorrect negative for comparison results in incorrect compensation values.

It is acceptable to use a combination of beads and cells to generate a compensation matrix, as long as you have matched positive and negative populations in each tube.

The use of cells for compensation particles is especially important for vital dyes, fluorescent proteins, reactive oxygen species, and other non antibody-based stains.

Compensation Rule 3: “Compensation controls must EXACTLY match the experimental fluorochrome and detector settings.”

Take the following 3 spectra: GFP, Brilliant Blue515™, and FITC. Each is excited by 488 nm light and measured in the “FITC” channel with a bandpass filter around 530/30 nm or so — however, their spectra are all subtly different (Figure 4).

Figure 4: Spectra of 3 different “green” fluorochromes.

Due to these differences, it is not possible to substitute one as a compensation control for the other.

Tandem dyes are manufactured and can degrade over time, so it is especially critical to use the identical tandem (same vial!) for setting compensation.

As shown in Figure 5, 2 different lots of the same fluorochrome (Cy7-PE) were acquired coupled to the same clone. Cells were stained with these 2 antibodies and the uncompensated data is shown. As can be seen from the different lines, if Lot #2 were used to compensate Lot #1, the resulting compensation values would be incorrect.

Figure 5: 2 different lots of the same tandem dye can have very different compensation values. Axes are labeled with excitation line (G=532 nm) and the band pass filter in front of the PMT.

Rule 3 also dictates that a single color compensation control must be collected for each fluorochrome in the panel. If 2 different panels are run at the same time, and these panels have different fluorochromes (especially tandem dyes), each panel needs its own compensation matrix.

Implicit in rule 3 is that the compensation control tubes must be treated identically to the samples. If you fix the cells, the controls must be fixed as well.

The effects of fixation or other treatments on fluorochromes is variable, so it is essential that the controls and samples are treated consistently.

A couple of final words regarding compensation. Compensation values are determined by a combination of fluorochrome properties and optimized instrument settings, not the carrier or antibody used.

When using beads as carriers, if the staining is off-scale, resist the temptation to decrease the voltage of the control, as this will only negatively impact your sensitivity. Rather, during the development of the assay, determine an appropriate amount of antibody with which to stain the beads, so that the fluorescent signal at the best voltage for the experiment meets the 3 rules above — especially that it is on-scale and within the linear dynamic range of the PMT.

What makes a good voltage? In practice, starting with an optimized voltage via peak-2 beads, CS&T, or other technique is a good start. Better still, is a voltration that takes into account the specific fluorochromes and cells being used in the experiment. How to go about determining this has been addressed here.

Finally, make sure to collect sufficient numbers of events. If using beads, at least 10,000 events should be collected. For cells, starting at a minimum of 30,000 events is good, but 50,000 is better. You want to have a sufficient number of events in your positive gate to have the best measure of the fluorescence.

So there they are, the 3 rules of compensation, and some important caveats that need to be remembered when setting up compensation controls for an experiment.

Looking beyond these 3 essential rules, make sure that the controls meet the other criteria addressed here, especially keeping the signal on scale and within the linear dynamic range of the detector. These steps will help ensure the highest quality compensation is obtained. Finally, avoid the temptation to manually adjust the compensation matrix, especially to make the data “look right”. Rather, determine what is causing the issues with the compensation by reviewing the data in the context of these rules. Also, make sure that a standard reference control is always run in the panel, to evaluate the whole staining process and help in troubleshooting when compensation “looks wrong”.

To learn more about the 3 Requirements For Accurate Flow Cytometry Compensation, and to get access to all of our advanced materials including 20 training videos, presentations, workbooks, and private group membership, get on the Flow Cytometry Mastery Class wait list.

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Tim Bushnell, PhD
Tim Bushnell, PhD

Tim Bushnell holds a PhD in Biology from the Rensselaer Polytechnic Institute. He is a co-founder of—and didactic mind behind—ExCyte, the world’s leading flow cytometry training company, which organization boasts a veritable library of in-the-lab resources on sequencing, microscopy, and related topics in the life sciences.

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