Top 5 Tips: Flow Cytometry

21st May 2026

Top 5 Tips for Flow Cytometry

Flow cytometry looks manageable until you're three hours into acquisition and realise your gating boundaries are off, your viability looks suspect, and one of your channels is bleeding into another. At that point, tracing the problem back is slow and frustrating. Most of these issues are decisions made before you ever put a sample on the cytometer. Panel design, fluorophore selection, controls — get these right at the start and the rest of the workflow is considerably more straightforward.

1. Know your instrument before designing your panel

This sounds like an obvious starting point. It's also one of the most common places panel design goes wrong, particularly when protocols are borrowed from another lab or adapted from a publication without checking instrument compatibility. Every cytometer has a fixed configuration of lasers, filters, and detectors. The fluorophores you choose need to match that configuration. A panel that works well on one instrument can perform poorly on another with a different laser line or filter set, even if the targets and antibodies are identical. Before you select a single fluorophore, map out what your instrument can actually detect. Know your laser lines, your detector bandpass filters, and which combinations create spectral overlap problems in your specific configuration. Panel design on paper means nothing if it doesn't match what's in front of you.

2. Pair fluorophore brightness to antigen expression

Not all fluorophores perform equally, and the mismatch between fluorophore brightness and antigen expression level is one of the more consistent sources of poor resolution in flow data. Bright fluorophores like PE and APC have high quantum yields and are best reserved for low-abundance or dimly expressed markers, where you need every photon to separate your positive population from the negative. Using a bright fluorophore on a highly expressed antigen doesn't improve your data and wastes a detector that could be doing more elsewhere in your panel. Dimmer dyes can handle highly expressed targets without issue. Matching brightness to abundance across your whole panel, rather than assigning fluorophores based on availability, will give you cleaner separation across every marker.

3. Always include a viability dye

Forward scatter and side scatter will not reliably exclude dead cells from your analysis. This is well established, and it's still one of the most common omissions in flow panels, particularly in labs new to the technique or running quick preliminary experiments. Dead cells bind antibodies non-specifically. If they're included in your analysis, they will skew your data in ways that are difficult to detect and easy to misinterpret. A population that looks real may simply be dead cells with non-specific labelling. A viability dye should be in every panel, every time, regardless of how quick the experiment is or how healthy your cells look. It's a small addition that removes a significant source of false positives.

4. Use FMO controls, not just isotypes

Isotype controls are widely used but have real limitations when it comes to setting accurate gating boundaries in multicolour panels. Fluorescence Minus One controls are more appropriate for most applications and give you considerably more accurate gating, particularly for dim or rare populations. An FMO control contains every fluorophore in your panel except the one you're gating on. This accounts for the spreading effect that multiple fluorophores have on the negative population in any given channel, which isotype controls don't capture. The result is a more accurate picture of where your negative population actually sits, and a gating boundary that reflects your real data rather than an artificial reference. For panels with more than two or three colours, FMOs are not optional if you want to gate with confidence.

5. Titrate every antibody in your panel

The manufacturer recommended concentration is a starting point. It is based on specific cells, specific staining conditions, and a specific instrument configuration. Change any of those and the optimal concentration shifts. Too high and you increase background, drive up non-specific signal, and reduce your staining index. Too low and your positive population loses resolution. Both outcomes make your data harder to interpret. Titrating each antibody to the concentration that maximises staining index while minimising background takes time upfront. It pays back in cleaner data, more reproducible results, and fewer experiments you have to repeat because the gating wasn't clean enough to draw conclusions from.

One more variable worth checking

Panel design and protocol optimisation can only take you so far if your antibodies aren't validated for flow cytometry. Clone selection, conjugate quality, and lot-to-lot consistency all affect how your panel performs. At St John's Laboratory, our flow cytometry validated range covers a broad selection of targets and conjugates, with application-specific data to support panel design from the start.

Explore the flow cytometry range →

Which of these five has caused you the most trouble in panel design or acquisition? Let us know in the comments.