Dr. Gareth Howell, Senior Experimental Officer and Flow Core Facility Manager at the University of Manchester answers your questions.
FMO, or fluorescence minus one, controls and isotype controls are different types of controls and aid the experiment in different ways. FMO controls allow you to determine how much spread is occurring between channels following compensation. They allow you to accurately determine where to gate for positive populations of cells by showing how much spread is on the negative population in the context of all the other colors in your experiment.
Isotype controls allow you to determine how much background binding may be occurring due to non-specific binding of the class-matched IgG, in particular to Fc receptors on cells such as monocytes and macrophages. Isotype controls should not be used in FMO controls. Be careful with isotype controls and remember a few points: 1) isotypes are IgG molecules so may show a degree of binding in the system you are using them in, 2) your isotype should carry the same fluorophore as your test antibody but, especially if you are using tandems, is unlikely to be of the same batch of fluorescent molecule complex so may be slightly different, and 3) ensure you are using the same concentration of isotype control as you are using for your test panel – use both at the same µg/mL concentration, not dilution as that is not sufficiently accurate.
FMO, or fluorescence minus one, controls are used to position gates during your cell analysis. As they contain all of the antibodies in your panel minus one, they allow you to determine how much spread you are getting in the negative population in the context of all of the other fluorochromes and post compensation. They are not factored into the design of the panel but knowing the signal spread matrix (SSM) is related to the FMO controls and is a key consideration in panel design.
First, check the technical aspect of the experiment – are the beads (comp beads) suitable for the species from which the antibody was isolated (not all comp beads are catch-all)?
If this is ok, check the antibody by staining comp beads with an alternative antibody but with a matching fluorochrome. When run at the same voltage do you get a good separation of positive and negative? If the fluorochrome is not a tandem conjugate you could use the alternative antibody as a surrogate (remember, compensation isn’t about the antibody, it’s about the fluorochrome). However, if it is a tandem then it’s advisable to keep using the same antibody as you use on your cells.
Check the voltages on that particular channel. If they have been set by an automated system they may not be ideal. Try performing a titration of the voltages (voltration) to determine a more optimal voltage setting. Increase the voltages in increments of 50 from 200 and watch for the appearance of the positive peak. Once the separation between positive and negative stops increasing then this is your optimal voltage. Optimal voltage should be set in this way using, for example, CD4 stained leukocytes. If this hasn’t addressed the problem, then you could try a titration of antibody on the comp beads. Beads do not need to be stained with the same dilution of antibody as your cells – they may need more or less antibody to give an appropriate signal to allow for accurate compensation to be performed.
There are a few options here and they may depend on the application and whether you are planning to fix the cells. If you are running unfixed samples then reagents such as 7AAD or Draq7 would work for you. These dyes emit in the far-red region of the spectrum, far away from the fluorescent proteins you are referring to. If you are planning to fix your cells, there are amine-reactive fixable viability dyes that emit in the far red, an example of this is the Zombie-NIR (Biolegend) and Proteintech’s Phantom Dye Red 780 Viability Dye. These are sensitive as there is little background from the autofluorescence of the live cells. They can also be used on live cells if you are planning to perform cell sorting, for example.
You should always titrate your antibodies when they are new to the panel, new to the system (e.g., a change in cell type), or if you have changed the fluorochrome. Titrations should be done on a 6- or 8-point scale starting from very low concentrations of antibody up to saturation of the antigen. Titrations should be done on the cells you are planning to stain later on. They should also be performed on a set number of cells that reflect the number of cells you plan to stain in your full experiment. Titrations can be performed in a two-stage strategy, whereby the lineage markers are titrated initially, then the secondary markers are titrated subsequently.
For more information on Flow Cytometry in general, please see our Complete Guide to Flow Cytometry.
The challenges of flow cytometry: an interview with David Lanham
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Aperture size is a measure of the width of the beam of light that is used to excite the cells in a sample during flow cytometry. A smaller aperture size results in a narrower beam of light, which can be more focused but may also result in less light reaching the cells. A larger aperture size results in a wider beam of light, which can cover more cells but may also result in the light being less focused.
There are several potential limitations of flow cytometry. One is that it can be difficult to accurately measure very small particles. Another is that the technique is not well suited for measuring particles that are not uniform in size or shape. Finally, flow cytometry can be expensive and time-consuming, so it may not be practical for all applications.
Flow cytometry is a powerful tool for analyzing blood samples because it can provide a great deal of information about the cells in the sample. This information can include the size of the cells, their shape, their surface markers, and their internal contents. This information can be very helpful in diagnosing diseases and in understanding how the cells in the blood are functioning.
A flow cytometer can be used to analyze the immune system in a number of ways. For example, it can be used to measure the number of cells in a given sample, the percentage of cells that are positive for a particular marker, the average size of cells, and the average granularity of cells.
Autofluorescence is when a fluorochrome (a molecule that absorbs and then re-emits light at a different wavelength) is present in a sample without being specifically bound to anything. In flow cytometry, autofluorescence can cause problems with data analysis because it can produce false positives – that is, events that appear to be positive for a particular marker but are actually just background noise. To avoid this, it is important to carefully control for autofluorescence when designing experiments and to use appropriate controls when analyzing data.
Check the voltages on that particular channel. If they have been set by an automated system they may not be ideal. Try performing a titration of the voltages (voltration) to determine a more optimal voltage setting. Increase the voltages in increments of 50 from 200 and watch for the appearance of the positive peak. Once the separation between positive and negative stops increasing then this is your optimal voltage. Optimal voltage should be set in this way using, for example, CD4 stained leukocytes. If this hasn’t addressed the problem, then you could try a titration of antibody on the comp beads. Beads do not need to be stained with the same dilution of antibody as your cells – they may need more or less antibody to give an appropriate signal to allow for accurate compensation to be performed.
Isotype controls allow you to determine how much background binding may be occurring due to non-specific binding of the class-matched IgG, in particular to Fc receptors on cells such as monocytes and macrophages. Isotype controls should not be used in FMO controls. Be careful with isotype controls and remember a few points: 1) isotypes are IgG molecules so may show a degree of binding in the system you are using them in, 2) your isotype should carry the same fluorophore as your test antibody but, especially if you are using tandems, is unlikely to be of the same batch of fluorescent molecule complex so may be slightly different, and 3) ensure you are using the same concentration of isotype control as you are using for your test panel – use both at the same µg/mL concentration, not dilution as that is not sufficiently accurate.
Dr. Gareth Howell, Senior Experimental Officer and Flow Core Facility Manager at the University of Manchester answers your questions.
You should always titrate your antibodies when they are new to the panel, new to the system (e.g., a change in cell type), or if you have changed the fluorochrome. Titrations should be done on a 6- or 8-point scale starting from very low concentrations of antibody up to saturation of the antigen. Titrations should be done on the cells you are planning to stain later on. They should also be performed on a set number of cells that reflect the number of cells you plan to stain in your full experiment. Titrations can be performed in a two-stage strategy, whereby the lineage markers are titrated initially, then the secondary markers are titrated subsequently.
First, check the technical aspect of the experiment – are the beads (comp beads) suitable for the species from which the antibody was isolated (not all comp beads are catch-all)?
Should you be using isotype controls?
The theory for using isotype controls is that the isotype control is an antibody with the same isotype as your target antibody but which binds to a non-target. This is supposed to be able to tell you what the non-specific binding is of your target antibody.
But, does it really tell you this? Are Isotype controls really acting as a control in your experiments?
Well, the first question you have to ask is are there primary targets for the isotype control on your cells of interest, and how do you know this?
For example, MOPC-173 is a common isotype mouse IgG2a kappa, it was first published in the 1970s has not been shown to bind any specific target. In fact, the production sheet from one vendor specifically states that this antibody was chosen as an isotype control after screening on a variety of resting, activated, live, and fixed mouse, rat, and human tissues.
Now, how much characterization was done as we continue to subset our populations of interest, we always find new and novel things. Look at the fact that murine reagent B220, thought to be on B cells of mouses has recently been shown to be present on a subset of human B cells as well.
So, looking at broad strokes, usually, because the isotype doesn’t bind, it’s not necessarily encouraging.
Second, do you know that the affinity of the variable region of your isotype control has the same affinity with the variable region of your target control? Now, how do you figure that out?
Third, is, of course, the fluorochrome to protein ratio.
Now, if you’re using PE, yes, the one-to-one is probably reasonable, but in many other cases, especially using things like Alexa Fluor or FITC, we don’t necessarily know what the F to P ratio is, unless you get it directly from the vendor, or you try to figure it out. While historically, isotype controls have been used to determine positivity, they really only help you show if blocking was effective.
I really discourage the use of isotype controls.
But you might be worried about what the reviewers of your publications will say.
This happened to a colleague who recently submitted a paper for publication, and one of the reviewers criticized the paper for the lack of isotype controls.
We put together a two-page document explaining why the isotype controls were not valid in this case and why they are not useful. We gave it to the investigator to send back to the reviewers. The paper ultimately got published.
If you are not convinced, read this paper by Anderson and co-workers from 2016.
In this paper, they do a really good job of showing you the value — or in this case the lack of value — of isotype controls.
FAQ
What are the 3 main components of flow cytometry?
What does flow cytometry tell you?
How many colors are in flow cytometry?
