As bright or brighter…

So the earlier posts were a lead-in to this one, which I hope will clarify this important issue in compensation.  When calculating compensation, it is important to make sure that your single stained control is as bright or brighter than your experimental sample for each color.  To illustrate why this is important, a GFP+ sample was measured in 6 detectors off of the blue laser:  525/50 to measure the GFP, and then also in 576/26 (PE), 610/20 (PE-TR), 675/20 (PE-Cy5), 695/40 (PE-Cy5.5), and 780/60 (PE-Cy7) to examine the spillover of GFP into these other detectors.  Populations are identified based on their GFP signal (neg, dim, mid, and bright), and plot titles refer to what was used to determine compensation from each channel (none=no compensation, then dim, mid, or high/bright indicates which population was compared to neg to calculate spillover).abob01

Compensation is basically determining the ratio of the two detectors in question: the signal in the primary detector, and then how much signal is detected in another channel (the spillover channel) for that intensity of staining.  This allows the researcher to subtract the spillover signal from other channels so that signal from that detector corresponds to that additional parameter.  In essence, the slope of the population’s signals is what is being determined:

 

abob02
If you are using a dim population, noise in the detection system reduces your sensitivity and so the signal will not be accurately measured.  When you look at a brighter population, it will probably not be compensated accurately.

abob03

Compensating with the dim population, the resulting means are neg 57, dim 57, mid 89, and bright 182.

In the case of a high spillover between channels, such as GFP spilling into the PE channel, it is possible to get decent compensation of the bright population using the “mid” population, since the signal is high enough to give you a more accurate measurement:

abob04
mean: 56, 52, 57, 49

But this may not be true for other spillovers in the same panel.  As you move further away from the optimal detector for that fluorochrome, only the brightest events will show spillover that can be detected above background:

abob05

In this case, using the “mid” population results in slight undercompensation of the brights (means of 72, 72, 73, 89) even though it “looks” ok.  (Of course, you should never use visualization to set compensation, but compare the means or medians of the negative and positive populations).

Even if different spillover values are generated using bright vs dim populations, the bright values will still work for the dim.  Changes in the spillover values have a much greater impact on events with high signal than dim.  For comparison, a 1% change in compensation in this example changes the mean signal of the dim by 5, mid by 24, or bright by 102 units in the spillover channel:

abob06

Correcting the spillover from 19.8% using the dim, which leads to the bright being undercompensated by 125 (73% off), to 21.02% using the bright makes the dim “overcompensated” by 4 which is hardly noticeable (5.5% off).

abob07

And just for completeness, below are tables of the spillover % for the different setups:abob08

…And the statistics tables for the means (none, dim, mid, bright)

abob09

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Compensation caution: matching beads

Other platforms may be different, but with BD’s Diva software you can only collect one “unstained” compensation control, so if it is necessary to mix what type of compensation particle you are using (cells for PI or DAPI, Arc beads for Live/Dead amine reactive dyes, rat comp bead or mouse comp bead for antibody from rat or mouse) you would have to define a different negative population for each type to get proper compensation.  In Diva, this is done by having an internal negative in the data file, and drawing a new interval gate on the histogram for that single stained control.  In the absence of this additional gate, the software refers to the “unstained control” tube for the comparison.
     At certain points in manufacturing, companies must change the batch of plastic they use for their bead, and the new batch can have different  fluorescence properties in some channels, especially in the 450nm emission range with 405 excitation (the “Pacific Blue” channel). If you mix your lot numbers of beads, you may get a mismatch. In this plot an older vial of “non-binding” bead is mixed with a newer vial (later expiration date) of unstained binding bead and you can clearly see the difference in intensity:
beadmismatch
When you add an antibody to this mix which is known to have no spillover into Pacific Blue (such as PerCP-Cy5.5) you still see a separation in the populations. The software is going to look at the positive population (green) and compare the mean signal for that population to the mean signal of the negative population (blue) in Pacific Blue, and since the positive is higher, this difference is attributed to spectral overlap and a compensation value is determined even though there is no fluorescence spillover, it is just a difference in the plastic.
compbead
If compensation is calculated in this case, any time you have PerCP-Cy5.5+ events, there will be signal inappropriately subtracted from the Pacific Blue channel.  On the left is the wrong compensation, on the right is compensation calculated with properly-matched beads.
comp mismatch

This was a 4 color experiement so the error was easy to notice, but in more complex panels this will not be the case, so make sure you do it right from the start!

I posted to on the Purdue message board about this a while back, here is the link towards the end of the thread with the previous messages attached, start at the bottom to go through the discussion chronologically.

https://lists.purdue.edu/pipermail/cytometry/2011-April/041298.html

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Full Disclosure: I did not receive any Compensation for this post…

In flow cytometry it seems the biggest source of confusion and angst is compensation.  As a first post, I’m going to re-publish a post I did elsewhere.

Compensation is somewhat tricky if you don’t understand the process. There are lots of good online tutorials and posts about compensation, but there are a few specific items I’d like to address and have “published” for viewing so I’ll throw my hat in the ring as well.

All fluorochromes will emit light across a range of wavelenghts, and if you are trying to take measurements for another fluor in a band that is included in this range, you will get background light (“spillover” or “spectral overlap”) from that other fluorochrome. To correct for this, most cytometry systems have some mechanism for “compensation” to remove this background. In essence, for any other parameter/detector  that you are measuring, you want your single positive population to have the same Mean Fluorescence Intensity (MFI) as the negative population so that any positive signal in the other channels can be attributed to that particular color, and is not spillover from something else. In the case shown, FITC signal is being detected in the PE channel, and once compensated has the same mean fluorescence in the PE channel as the unstained.

Many companies now offer antibody-capture bead kits for compensation (Invitrogen, BD, Bangs Labs, Spherotech) and they usually include a “binding bead” and “non binding bead” in the same box. For example, BD offers “Compbeads” which bind to a specific species’ IgG (mouse, rat, hamster), so for a mouse-anti-Human CD4 antibody, you’d need the mouse binding bead, but for a rat-anti-mouse CD19 you’d need the rat binding bead. For Invitrogen’s ARC beads which are used to compensate their amine-reactive Live/Dead fixable stains, the binding beads are coated with amine groups so the reactive dye will bind to the bead. The Live/Dead dye will NOT bind to antibody-capture beads.

In order to correctly do this calculation, it is critical that you are comparing particles of the same type– if you are using stained cells, you want to compare the signals to the *same type* of unstained cells. If you are using beads, you want to compare to the exact same unstained bead. The “non-binding” bead is included in the kits for this purpose-they are the same bead as the binding one in the kit, but will not stain with the reagent and thus remain negative. You can include them both in the same tube, although you should wash the beads after staining to remove free dye if you do so.

Mismatches in your positive and negative particles will result in compensation errors, which could dramatically impact your data. More on that later…  Mike

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To New England Cytometry’s new website!

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