5 Common Flow Cytometry Questions, Answered

I want to thank all of you who send us your questions about flow cytometry, so I thought I would dip into the old email bag and answer a few of the common ones here.  If your question isn’t answered this time, look for it to be answered in a future blog post.  Of course, if you want us to cover a specific topic, drop us a line. 

1. How Fast Can I Go?

This is  a common question. The allure of the ‘hi’ button is hard to resist.  The faster you go, the sooner you are finished with data acquisition, and if you run samples in a shared resource facility, time is money.   

Speed is not everything, and in fact, can be detrimental to your data acquisition.

The rate that a flow cytometer runs is set by the sheath fluid pressure.  Since we do not adjust this, the way we increase our event rate is to change the differential pressure between the sheath fluid and the sample.  The bigger the difference, the more events can pass through the laser intercept. 

While increasing the differential pressure increases the number of events per second, it also negatively impacts the data in several ways:  

• Increase in coincident events, which are removed as doublets
• Increase in abort rate, meaning lost data
• Increase in the CV of the data, compromising the sensitivity of the assay

Thus, while you are getting more total events, you are losing them as aborts and conicident events.  More importantly, the sensitivity is compromised.

Shown in Figure 2 are the results of changing the flow rate on a cell cycle experiment.  When running a cell cycle assay, the quality of the data is judged by the CV of the G0/G1 peak.  In the figure, the cells were stained and run at different flow rates.  The yellow lines are there to orient to the G0/G1 peak.  If you compare the low to the medium, there is an increase in the spreading of the data, and this is especially pronounced in the G2/M peak, as well as the S phase.  Moving to high it is clear that the G0/G1 peak is very compromised and its almost impossible to identify the S page in this data. 

Best practices is to run your samples at low flow rate.  Even more important is to not run one sample at low flow rate and a second one at high flow rate.  If you were to use the low flow rate sample to set your gating strategy, you would lose data from the high flow rate sample due to this spreading of the CV. 

One way this has been solved is using acoustic focusing, which has been commercialized in the Attune flow cytometer from ThermoFisher. In this system, the cells are initially hydrodynamically focused like other flow cytometers.  The trick is that a standing acoustic wave is generated that pushes the cells to the center of the core stream, so even when running at high flow rates, the cells are aligned, which reduces the impact on the CV of the data compared to non-acoustic focused instruments.

2. What’s All The Buzz About The Fluorescence Minus One Control?

Gating is a data reduction tool.  When we place a gate on a plot, we are making a choice of what cells we will include and which ones are excluded.  Therefore, getting the position of these gates is important so the wrong cells are not carried forward for downstream analysis.  To do this, it is important to have the correct controls, which will allow you to properly position the gate. 

Historically, something called the isotype control was used as one of the tools to set positivity.  There are issues with this control, which I have written about in the past, and I direct your attention to that article.

Enter the Fluorescent Minus One control.  This is a tube where the cells are stained with all the fluorochromes in the panel but one.  This helps to identify the upper limit of your background signal, based on the contribution of the other fluorochromes into the channel of interest.  An example is shown below.

On the left is the completely unstained sample, on the right is the fully stained sample.  The middle panel is the FMO control, which contains all the fluorochromes except PE.  If the positive gate was set using the red (Unstained) line, it is clear from the middle plot that gate would call the cells above that red line PE positive, which they cannot be since there is no PE in that tube.  Thus the FMO helps to identify where the positive gate should be set, which is the blue line.  

The FMO control is especially valuable in rare event analysis and for emergent markers. It should be used anywhere that identification of the target cells is important.  

3. What Is A Good Cutoff To Say My Population Is Positive?

In other words, what is the appropriate number to call my population a real population? Is it 1%, 5%?  This is especially of interest in rare event analysis.  How many cells do I need to collect to convince my reviewers that the population is real.   

This is a great question that was discussed in an article by Mario Roederer.  In this article, points out that:

“The question of whether events are “real” or not is fundamentally inappropriate. Of course they are “real”. The appropriate question is: do the events represent what the researcher claims they are…”

He goes on to further conclude:

“In conclusion, there is no theoretical reason to employ an artificial threshold number of events, below which a frequency is deemed “negative”. The assessment of “positivity” can only be made by comparison of the measurement against a set of control samples, using standard statistical tools to compare the frequencies.”

Take a look at this data.  On the left are the results of analysing ten control samples, and the right ten experimental samples.  The data is plotted and a two way t-test was performed, with a p=0.05. As is shown, there is significant difference between the two populations.  

4. What Should I Use For Compensation Controls – Beads Or Cells?

It is important to remember that compensation is a property of the fluorochrome, not the carrier.  The carrier serves to get the fluorochrome to the interrogation point for measurement.  So antibody capture beads or cells are both useful controls.  So which one should you choose?  

In 2011, FlowJo published a post on the three rules of compensation, which you can read here. The second rule of compensation says, “Background fluorescence should be the same for the positive and negative control.” 

When we compensate, were are comparing the signal in the secondary channel (fig 5: blue box), and calculating the value that will set the mean of this population equal to the mean of the negative (fig5: purple). 

Cells are useful controls because you will be labeling the cells of interest, can address issues of autofluorescence and don’t cost anything.  Beads are useful because they can bind the exact antibody/fluorochrome (compensation ruler #3), they generally provide clear separation between the positive and negative, and are useful to not sacrifice the cells on a control.  What is especially important is to not mix beads and cells for the same channel.  For example, many systems allow for the use of a ‘universal’ negative to use for calculating compensation.  This is something to avoid as showing in figure 6.

In figure 6, unstained cells are indicated in green.  The positive signal is on beads.  If one draws a line from the mean of the negative population to the mean of the positive signal, you can see how a mismatch can impact compensation.  

The best practice is to have a negative and a positive in each compensation control. In this way, it is easy to use cells for those fluorchromes you need to (viablity dyes, redox dyes, etc), while using beads for the other fluorochromes.  

One caveat for this is in spectral cytometry.  There are occasions that cells are better for performing spectral deconvolutions, and this has to be determined empirically.

In general, reusing the compensation matrix can violate the 3rd rule of compensation which states “Your compensation control must be matched to your experimental control.”  Embedded in this rule is that you need to have the identical fluorochrome and identical instrument sensitivity. 

5. Can I Reuse My Compensation Matrix?

In general, reusing the compensation matrix can violate the 3rd rule of compensation which states “Your compensation control must be matched to your experimental control.”  Embedded in this rule is that you need to have the identical fluorochrome and identical instrument sensitivity. 

Tandem dyes rely on Förester resonance energy transfer (FRET) between the donor and acceptor fluorochromes.  This process is heavily dependent on the distance between the donor and acceptor, and the efficiencies of transfer is inversely proportional to the 6th power of the distance between the donor and acceptor.  Tandem dyes can be disrupted by oxidation, temperature changes and certain fixatives.  If your staining mix has tandem dyes, there can be changes over time, which impact the compensation, so using a matrix from an earlier time point may result in incorrect compensation for the tandem when used later on. 

Instrument sensitivity is also important, and this can be measured in several ways.  There are several ways that instrument sensitivity can change such as: after a realignment, replacement of  an optical element (detector, laser, filter), cleanliness of the machine and more.  While beads can be use to help minimize this, unless the researcher started using beads at the beginning of the experiment, beads won’t help. 

Thus it is easiest and the best practice to run compensation every time.  Now, if the researcher has not remembered to prepare compensation tubes, this paper by Andersen and coworkers offer some suggestions on reusing a compensation matrix.   

So there you have it, common questions answered.  Hopefully this gives you some more insight into ways to improve your own work.  Ultimately understanding the theory of how the flow cytometer works is critical to generating high quality, reproducible date.  Until next time, keep those questions coming and may your stream always be laminar.

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