A major advantage of flow cytometry over other immunoassay techniques is that it allows researchers to analyze extremely high numbers of single cells relatively quickly. In recent years, it has become common practice to detect a target of interest alongside multiple cell markers, with experiments routinely designed to measure 10–20 parameters in parallel. However, choosing the right antibodies for flow cytometry is fundamental to experimental success. Here, we look at some of the challenges of antibody selection for flow cytometry and provide tips for deciding which reagents are best suited to prove a particular hypothesis.
Flow cytometry data is susceptible to misinterpretation
“In most immunostaining assays, researchers are provided with information about antibody performance beyond simply the amount of antibody binding to the sample,” notes Christopher Manning, associate director of flow cytometry and high content analysis at Cell Signaling Technology. “Examples include the molecular weight of bound proteins (western blot), the localization of binding (immunohistochemistry and immunocytochemistry), or the orthogonal confirmation of target specificity using a second antibody (sandwich ELISA). In contrast, when using a traditional flow cytometer, fluorescence intensity is the only indicator of antibody performance. As a result, it can be difficult to know when an antibody is exhibiting non-specific binding. This makes flow cytometry data more susceptible to misinterpretation than other antibody-based assays.”
Fluorophore selection compounds the potential for error
With many flow cytometry experiments designed for multiplexing, a further challenge of antibody selection relates to panel design. It is critical that antibodies that will be used simultaneously are conjugated to fluorophores with minimal spectral overlap, yet it is not always the case that the chosen antibody clone is available bound to a fluorophore that fits with the existing panel. “It is recommended to pair highly expressed markers with dim fluorophores, and vice versa,” explains Dr. Alexandra Wittmann, senior scientist at Abcam. “It is also good practice to distribute dyes with overlapping spectra on different cells rather than using them to bind multiple targets on the same cell. Where an appropriate antibody-conjugate is unavailable, it is now possible using commercial kits to rapidly label antibodies with fluorophores in-house.”
Assess the extent of supplier validation
A good place to begin when selecting antibodies for flow cytometry is to assess the extent of validation performed by the supplier. First and foremost, the antibody should be proven to work in flow. “Ideally, validation testing will have been performed using appropriate positive and negative controls,” says Wittmann. “These might be cells from a primary source containing various cell types with differential target expression, or knockout cell lines that offer a true negative control.” Manning adds that supporting data should include validation across complementary assays and against isotype controls to confirm antibody specificity, and evaluation of the sensitivity of the antibody. He also notes that non-specific cross-reactivity is less problematic when antibodies are used on live cells than when the whole intracellular landscape is available for binding.
Image: Overlay histogram shows HAP1 wildtype (green line) and HAP1-MKI67 knockout cells (red line) stained with an anti-Ki67 recombinant rabbit monoclonal antibody (ab16667), followed by an anti-rabbit IgG H&L secondary antibody (AlexaFluor® 488, ab150081). Image provided by Abcam.
“Obviously, suppliers cannot perform every single validation step imaginable,” says Dr. Mona Al-Maarri, global product manager for flow cytometry reagents at Miltenyi Biotec. “But knowing which validation steps have been carried out allows you to decide which, if any, further validation is required before you begin your experiment. This not only helps ensure the success of your experiment, but saves time and money, and can make a huge difference to the progress of your research.” She also suggests bearing in mind that the most-cited antibodies are not necessarily the best performers—they may have just been available the longest.
Use suppliers’ recommendations and protocols as guidelines
Antibody datasheets are a valuable source of information, containing details of the antibody clonality and isotype, species reactivity, target epitope location (surface or intracellular), and tested applications. They also typically include recommended use conditions, namely the protocol that was employed for antibody validation and a suggested antibody concentration range for each tested application. This provides researchers with a strong starting point from which to set up their own protocols.
“A good antibody datasheet should show testing data proving that the antibody is suitable for the chosen application,” reports Wittmann. “In a best-case scenario, the supplier will have used cells that are relative to the system used by the researcher and will have shared any additional findings such as whether fixation prior to staining impacts epitope recognition or whether certain permeabilization agents denature surface-bound fluorophores. But even in the unlikely scenario that an antibody suppliers’ validation process exactly mirrors the intended use, it is essential that researchers test antibodies in their own experimental system. This allows for identification of the ideal antibody concentration and for fine-tuning the staining protocol for the cell type of interest.”
One essential test that should be carried out involves titrating the antibody across known positive and negative conditions to find the optimal dynamic range, where signal in the positive condition is maximized while little or no signal is present in the negative condition. It is also wise to repeat this titration each time a new lot of antibody is received to ensure consistency and comparable experimental results. Although it is widely recognized that different antibody lots can vary in terms of performance, the conjugated fluorophore is another variable that must be considered.
Antibody suppliers are here to help
It can be argued that the reproducibility crisis has caused researchers to focus greater attention on the antibodies they use and to demand increasingly rigorous validation efforts from antibody suppliers. “Speaking from personal experience, many researchers have historically relied on antibodies and workflows established in their laboratories without knowing if antibody reagents have been thoroughly validated for the experimental setup or not,” says Al-Maarri. “This mindset is now changing and antibody suppliers are supporting this by providing ever more information on product datasheets regarding antibody specificity, sensitivity, and reproducibility.”
She adds that while the potential for spill over and the lack of spatial context are inherent drawbacks of flow cytometry, the opportunity to stain around 10–20 antigens simultaneously and to analyze high numbers of single cells within reasonable timeframes more than compensates for this. “With careful experimental set up—in particular, ensuring a well-matched combination of conjugated antibodies and thorough compensation—flow cytometry can identify even rare sub-populations for deeper insights into target cells’ nature and complexity,” she says. “Furthermore, dead cells can easily be excluded and there is no bleaching effect to worry about, meaning flow cytometry remains one of the most widely used immunoassay techniques available.”