Technological advancements in fluorescence flow cytometry and an ever-expanding understanding of the complexity of the immune system have led to the development of large flow cytometry panels reaching up to 43 colors at the single-cell level. However, as panel size and complexity increase, so too does the detail involved in designing and optimizing successful high-quality panels fit for downstream high-dimensional data analysis. In contrast to conventional flow cytometers, full-spectrum flow cytometers measure the entire emission spectrum of each fluorophore across all lasers. This allows for fluorophores with very similar emission maxima but unique overall spectral fingerprints to be used in conjunction, enabling relatively straightforward design of larger panels. Although a protocol for best practices in full-spectrum flow cytometry panel design has been published, there is still a knowledge gap in going from the theoretically designed panel to the necessary steps required for panel optimization. Here, we aim to guide users through the theory of optimizing a high-dimensional full-spectrum flow cytometry panel for immunophenotyping using comprehensive step-by-step protocols. These protocols can also be used to troubleshoot panels when issues arise. A practical application of this approach is exemplified with a 24-color panel designed for identification of conventional T-cell subsets in human peripheral blood. © 2021 Malaghan Institute of Medical Research, Cytek Biosciences. Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1 : Preparation and evaluation of optimal spectral reference controls

Support Protocol 1 : Antibody titration

Support Protocol 2 : Changing instrument settings

Basic Protocol 2 : Unmixing evaluation of fully stained sample

Basic Protocol 3 : Evaluation of marker resolution

Support Protocol 3 : Managing heterogeneous autofluorescence

Basic Protocol 4 : Assessment of data quality using expert gating and dimensionality reduction algorithms

Technological advancements in fluorescence flow cytometry and an ever-expanding understanding of the complexity of the immune system have led to the development of large panels reaching 40 fluorophores in the Optimized Multicolor Immunofluorescence Panel (OMIP; Park, Lannigan, & Jaimes, 2020) and 43 colors in a technical note (Sahir, Mateo, Steinhoff, & Siveen, 2020). In contrast to conventional flow cytometry, which primarily measures the peak emission of each fluorophore in a target detector, full-spectrum flow cytometry uses a larger number of detectors with narrow band-pass filters. This allows the entire emission spectrum for every fluorophore to be captured across all laser lines, creating a detailed signature of each fluorophore. This makes it possible to distinguish fluorophores with very similar emission maxima but unique overall spectral fingerprints, increasing the flexibility in fluorophore selection. This feature, coupled with an instrument designed to maximize the detection of emitted light and highly efficient avalanche photodiodes, provides improved detection efficiencies that translate to better detection limits and higher signal resolution (Feher et al., 2016). While these features, unique to full-spectrum flow cytometry, provide the high-quality signals and low noise needed for successful high-dimensional panels, many of the same panel design considerations from conventional flow cytometry still apply. Common characteristics, together with those specific to full-spectrum flow cytometry, have been previously described (Ferrer-Font, Pellefigues, et al., 2020; Park et al., 2020). Now that hardware limitations hindering the use of highly overlapping dyes have been overcome, the main limitations for successful large panel design are the spillover-spreading error inherent to the use of fluorescence and the number of fluorophores available with unique spectral signatures. Many companies have recently begun developing spectrally distinct fluorophores, thus advancing the number of markers available to be analyzed in a single experiment.

Increasing the number of markers in a panel consequently increases the probability of compromising the resolution of the markers and populations of interest. The theoretical approach to panel design aims to avoid issues would that prevent resolution of every marker in the panel; however, in practice it is challenging to perfectly predict the impact of co-expression, variations in marker expression levels, and the performance of each specific reagent. Furthermore, it is difficult to anticipate the accuracy of reference controls for optimal unmixing results. For example, certain fluorophores emit slightly different spectra when bound to compensation beads or in the presence of different buffers.

One obstacle that has always impacted panel design and performance is the unique autofluorescence (AF) signatures of different sample and cell types. Cellular AF levels can vary depending on the type and metabolic state of cells (Mayeno, Hamann, & Gleich, 1992; Roederer, 2016; Shi et al., 2017) as well as sample preparation and staining procedures. This translates into different AF brightness levels and distinct spectral signatures in the samples being analyzed. Full-spectrum flow cytometry can resolve cellular AF signatures and ensure that they are not attributed to any of the fluorophores used. This can improve the signal-to-noise ratio and resolution of markers attached to fluorophores that emit closest to AF maxima (Ferrer-Font, Pellefigues, et al., 2020) in highly autofluorescent tissues such as brain, lung, skin, intestine, and tumor (Schmutz, Valente, Cumano, & Novault, 2016). It is therefore highly recommended to characterize the AF spectrum of an unstained sample from the tissue or cell type of interest prior to panel design. This will provide useful information during fluorophore selection, ensuring that fluorophores are not allocated to areas of the spectrum where AF dominates.

Once a high-dimensional full-spectrum flow cytometry panel has been optimally designed (Ferrer-Font, Pellefigues, et al., 2020), the step-by-step protocols presented here provide a series of practical steps for full and successful optimization. As outlined in Figure 1, the main procedures include evaluation of spectral reference controls (SRCs; see Basic Protocol 1), evaluation of unmixing of the fully stained (FS) sample (see Basic Protocol 2), evaluation of marker resolution (see Basic Protocol 3), and assessment of data quality (see Basic Protocol 4). Before any of these methods are performed, it is essential to titrate the antibodies used as well as any viability dye used. This is described in Support Protocol 1. An additional protocol describes changes that can be made to the instrument settings (see Support Protocol 2). Methods for evaluating and mitigating autofluorescence are included in Basic Protocol 3 and Support Protocol 3. Together, these protocols can also be used when troubleshooting a panel to identify sources of problems and provide insights into fixing them.

A 24-color panel optimization for identification of conventional T-cell subsets in human peripheral blood is provided to illustrate these procedures. The protocols were developed using the five-laser (5L) Cytek Aurora (Cytek Biosciences), but should be adaptable to any spectral flow cytometer. The protocols were designed for new full-spectrum flow cytometry users. Once familiarity and experience with specific tissue types is achieved, it is possible to modify the steps and order of the protocols to reduce the overall time spent evaluating the panel. If a modified approach is taken, it is recommended that all the overall goals of the protocols (as outlined in Fig. 1) still be carried out.

This protocol is divided in two sections. The first is for preparation of SRCs and ensures generation of the high-quality controls required for accurate unmixing. The steps describe the staining of polystyrene compensation beads and cryopreserved PBMCs with surface-labeling antibodies. They can be adapted for other tissues or staining procedures such as intracellular staining. It is important to mention that the preparation procedure will also guide users to prepare a fully stained (FS) sample and fluorescence minus one (FMO) controls as the protocols are very similar; these will be used later for evaluating the unmixing of the FS sample (see Basic Protocol 2). If preferred, FS and FMO samples can be stained separately, but treatment of samples should be kept identical. Importantly, to successfully complete this protocol, SS and FS cells should also be treated the same (antibody concentration, incubation time/temperature, fixed/unfixed, etc.).

The second section of the protocol is for evaluation. This aims first to check the quality of the acquired SRCs and then to evaluate whether there are any spectral mismatches between beads and cells for each fluorophore, which is accomplished by assessing how well the beads unmix the cells using N × N plots for all markers. Unmixing accuracy must be assessed on the actual sample (cells) to be used in the assay. In some cases beads will be acceptable as SRCs and in other cases it may be necessary to use cells. When this protocol is completed, the optimum SRCs (cells or beads) will be determined for future unmixing. If desired, well-characterized and high-quality controls can be stored for future use.

Materials

• Cryopreserved PBMCs

• Phosphate-buffered saline (PBS; Gibco, cat. no. 14190-250)

• Fetal bovine serum (FBS; Gibco, cat. no. 10091-148)

• Polystyrene compensation beads (e.g., UltraComp eBeads, Life Technologies, cat. no. 01-222-42)

• FACS staining buffer: PBS with 2% bovine serum albumin (BSA; MP Biochemicals, CAS no. 9048-46-8) and 0.2% sodium azide (Sigma-Aldrich, cat. no. S8032)

• Zombie NIR Fixable Viability Kit (BioLegend, cat. no. 423106)

• Human TruStain FcX (Fc Receptor Blocking Solution; BioLegend, cat. no. 422301)

• Antibodies (see Table 1 for list; see Support Protocol 1 for titration)

• Brilliant Stain Buffer Plus (BD Biosciences, cat. no. 566385)

• 37°C water bath (e.g., Julabo Ecotemp TW12)

• Hemocytometer (e.g., Hawksley Counting Chamber) and coverslips

• 96-well U-bottom plate (In Vitro Technologies, cat. no. 353077)

• Filters with 0.65-μm or smaller pore size (optional)

• Spectral cytometer (e.g., Cytek Aurora)

• 5-ml polypropylene round-bottom flow tubes (In Vitro Technologies, cat. no. 352008)

• Cytek SpectroFlo software

• Data analysis software for analyzing FCS files (e.g., FlowJo or FCS Express)

Prepare and acquire samples

1.Thaw PBMCs quickly in a 37°C water bath and add to 5 ml PBS with 2% FBS.

2.Centrifuge 5 min at 500 × g , room temperature, and carefully flick off the supernatant.

3.Resuspend cells in PBS with 2% FBS and count viable cells on a hemocytometer.

4.Centrifuge as before and carefully flick off the supernatant.

5.Resuspend cells in PBS with 2% FBS to a concentration of 5 × 10⁶ viable cells/ml.

6.Distribute 100 μl suspension per well of a 96-well U-bottom plate, allocating one well per fluorophore in the panel and one well for an unstained sample.

7.Add one drop of vortexed compensation beads plus 150 μl FACs staining buffer to a second set of wells, allocating one well per fluorophore in the panel except the viability dye.

8.Centrifuge plate 5 min at 500 × g , 4°C, and flick off the supernatant.

9.Prepare viability stain in PBS according to the titration result (see Support Protocol 1) and add 100 μl to the live/dead cell control and FS sample, if included. Resuspend remaining wells with 100 μl PBS. Incubate for 15-30 min at room temperature, protected from light.

10.Centrifuge as above and flick off the supernatant.

11.Block Fc receptors of the cell samples using 100 μl of 1:40 Fc block in FACS staining buffer. Add an equal volume of FACS staining buffer to the beads. Incubate for 10 min at 4°C.

12.Centrifuge as above and flick off the supernatant.

13.Remove aggregates from antibody stocks by centrifuging 5 min at 16,000-18,000 × g , 4°C (Aass et al., 2011; Ayers et al., 2011; van der Vlist, Nolte-’t Hoen, Stoorvogel, Arkesteijn, & Wauben, 2012) and/or by filtering using a pore size of 0.1-0.65 μm (Inglis et al., 2015).

14.Prepare SS, FS, and FMO control antibody mixes by diluting stocks in FACS staining buffer according to titration results (see Support Protocol 1). Be sure to pipette from the top of the liquid to avoid centrifuged aggregates.

15.Stain all cells and beads with 100 μl of the appropriate antibody mix and incubate in the dark using the incubation time and temperature that will be used in the final assay.

16.Centrifuge as above and flick off the supernatant.

17.Wash twice with 200 μl FACS staining buffer, centrifuging again after each wash.

18.Resuspend in 200 μl FACS staining buffer.

19.Acquire on a Cytek Aurora, taking care to meet the following acquisition criteria:

0. Cytek assay settings are used as a starting point for instrument setup.

1. The scatter profiles of cells and beads are on scale and the FSC area scaling factor (ASF) is optimized (see example in Fig.2B).

2. All fluorescence signals are on scale (<4 × 10⁶). This can be assessed in the full-spectrum plot or in individual plots for every detector.

3. All tubes are recorded with the same fluorescence gain settings for each detector.

4.         Sufficient events are recorded to find a clear positive signal.

        A minimum of 300 events is needed for each positive and negative population. A good starting point is 5000 total events for beads and 30,000-50,000 total events for cell controls, although it may be necessary to record >50,000 cells to get at least 300 positive events of similar fluorescence intensity for rare markers.

Evaluate results (see Video 1)

20.In the SpectroFlo software, check the raw reference control data and verify that the acquisition criteria in the previous steps have been met:

0. The scatter profiles of the cells and beads are on-scale, clean, and easily gated, and the FSC ASF has been optimized (Fig.2B).

1. Cytek assay settings or a close alteration (also see Critical Parameters discussion of Complex Samples and see Support Protocol2) have been used, and all tubes have been recorded with the same settings.

Antibody titration is the crucial first step in developing high-dimensional flow cytometry panels. Using the incorrect antibody concentration can increase spread, decrease resolution, increase aggregation of reagents, and give rise to nonspecific binding (Stewart & Stewart, 1997), all of which result in poor panel performance and/or inaccurate results. Ideally, titrations should be carried out in the tissue that will be used in the assay. This is not always feasible, however, if the tissue in question is rare or difficult to work with, or the cells of interest are found at low frequencies within the tissue. In such cases, it is suggested to first add a lineage marker to the mix to help identify the cells that express the rarer marker or to use a surrogate tissue in which the marker is more abundant and simpler to process. Results from this type of titration should always be validated using the tissue of interest.

It is important that readout functional markers are titrated under the maximally activated conditions that will be used in the assay. Staining conditions should also be identical between titration and experiment to prevent spectral pattern mismatches and poor unmixing results.

This protocol describes the preparation of cryopreserved peripheral blood mononuclear cells (PBMCs) for titration of extracellular antibodies. It can be easily adapted and used as a general guideline for antibody titration using other tissues or staining procedures.

Materials

• Cryopreserved PBMCs

• Phosphate-buffered saline (PBS; Gibco, cat. no. 14190-250)

• Fetal bovine serum (FBS; Gibco, cat. no. 10091-148)

• Zombie NIR Fixable Viability Kit (BioLegend, cat. no. 423106)

• Human TruStain FcX (Fc Receptor Blocking Solution; BioLegend, cat. no. 422301)

• Polystyrene compensation beads (e.g., UltraComp eBeads, Life Technologies, cat. no. 01-222-42)

• Antibodies (see Table 1)

• FACS staining buffer: PBS with 2% bovine serum albumin (BSA; MP Biochemicals, CAS no. 9048-46-8) and 0.2% sodium azide (Sigma-Aldrich, cat. no. S8032)

• 37°C water bath (e.g., Julabo Ecotemp TW12)

• Hemocytometer (e.g., Hawksley Counting Chamber) and coverslips

• 96-well U-bottom plate (In Vitro Technologies, cat. no. 353077)

• Filters with 0.65-μm or smaller pore size (optional)

• Spectral cytometer (e.g., Cytek Aurora)

• 5-ml polypropylene round-bottom flow tubes (In Vitro Technologies, cat. no. 352008)

• Data analysis software for analyzing FCS files (e.g., FlowJo or FCS Express)

Prepare and acquire samples

1.Thaw PBMCs quickly in a 37°C water bath and add to 5 ml PBS with 2% FBS.

2.Centrifuge 5 min at 500 × g , room temperature, and carefully flick off the supernatant.

3.Resuspend cells in PBS with 2% FBS and count on a hemocytometer.

4.Centrifuge as before and carefully flick off the supernatant.

5.Resuspend cells in PBS with 2% FBS to a concentration of 5 × 10⁶ cells/ml.

6.Distribute 100 μl suspension per well of a 96-well U-bottom plate, allocating six wells to each antibody being titrated. Include additional wells for unstained and live/dead controls (see example plate plan in Fig. 3A).

7.Centrifuge plate 5 min at 500 × g , 4°C, and flick off the supernatant.

8.Stain titration samples and live/dead controls with a viability dye that will not cause significant spillover into the fluorophore being titrated.

9.Centrifuge as above and flick off the supernatant.

10.Block Fc receptors by applying 100 μl of a 1:40 dilution of Human TruStain FcX and incubating 10 min at 4°C.

11.Add one drop of vortexed compensation beads per well for the appropriate single stain (SS) controls.

12.Centrifuge as above and flick off the supernatant.

13.Remove aggregates from antibody stocks by centrifuging the vials 5 min at 16,000-18,000 × g , 4°C, or by filtering using a pore size of 0.65 μm or smaller.

14.Create a dilution series for each antibody being titrated by diluting stock solutions in FACS staining buffer. Be sure to pipette from the top of the liquid to avoid the spun-down aggregates.

15.Add 100 μl of each antibody dilution to the corresponding sample in the plate. Add FACS staining buffer to unstained and live/dead controls. For compensation beads, use 100 μl of the 1:100 dilution.

16.Incubate using the time and temperature that will be used in the final assay.

17.Centrifuge as above and flick off the supernatant.

18.Wash twice with 200 μl FACS staining buffer.

19.Resuspend in 200 μl FACS staining buffer and acquire on a Cytek Aurora using Cytek assay settings.

Analyze data

20.Open titration samples in a data analysis software used for analyzing FCS files.

21.Generate median fluorescence intensity (MdFI) values for the positive and negative populations for each concentration.

22.Generate standard deviation (SD) values for the negative population for each concentration.

23.Calculate the stain index (SI) using the equation SI = (MdnFI pos – MdnFI neg) / (2 × SD neg).

24.Select the concentration that gives rise to the best stain index without giving rise to a positive shift of the negative population.

In some cases an adjustment of fluorescent gain settings away from the optimized settings cannot be avoided, for instance, if a bright reporter protein (e.g., eGFP) is off-scale. Such adjustments must be carried out carefully to minimize impact on all spectral signatures within the panel. Adjustments can be made detector by detector or to a whole detector array at one time, where all detectors of a given laser line are changed simultaneously. Reduction of the whole array is the recommended approach, as manual alteration of single detectors is prone to distortion of spectral signatures, where the ratio of brightness from detector to detector is not maintained. Regardless, reducing whole detector arrays can lead to peak emissions of spectral signatures being shifted to an incorrect detector (potentially on a different laser line). Therefore, all changes must be checked to ensure that signatures remain as expected based on Cytek's Full Spectrum Viewer, published fluorophore guidelines, or historical data.

1.Observe which fluorescent channel(s) has an off-scale signal.

2.Use trial and error to determine the gain setting that is appropriate to bring the emission peak on-scale (i.e., <4 × 10⁶ on the Aurora).

3.Reduce secondary detector arrays with off-scale signals by the same percentage as the reduction made in step 2.

4.Use Cytek's Full Spectrum Viewer, published fluorophore guidelines, or historical data to check that the altered spectral signature retains the correct overall pattern and that the primary emission peak has not been reduced below any secondary emission peaks.

5.If adjustments to other laser lines are required, continue reducing the respective detector arrays by the same percentage as in step 2.

6.Repeat steps 4 and 5 until the spectral signature appears as expected or all arrays have been reduced by the same percentage.

7.Record all samples at the adjusted settings.

Before any analysis can be undertaken, the FS sample must be checked to ensure that clean data can be obtained through removal of artefacts such as doublets, dead cells, and aggregates. It is also necessary to verify that there is positive staining for all markers in the panel, taking into consideration the biology of each marker. Once this has been asserted, it must be determined whether the SRCs selected (either beads or cells) successfully unmix the SS cells in Basic Protocol 1 can also be used to successfully unmix the FS sample (Video 2).

Once the best possible unmixing has been achieved using Basic Protocols 1 and 2, the resolution of each marker in the FS sample must be compared to the SS cell controls. Assessing the spread of the negative population and/or shifts in the positive signal will provide an indication of whether there is any loss of resolution of markers in the panel when fully stained. This assessment is best achieved by overlaying each marker in the SS cell sample onto the FS sample. If any reduction in resolution is seen, the impact of this can be further investigated to identify if it will impact the ability to identify populations of interest using an established gating strategy. Ideally, the same number of cells from the same tissue type should be stained and acquired for all samples. In practice, however, this is not often feasible, and downsampling can be used to achieve identical cell numbers across samples during analysis. In theory, the only difference between the SS and FS cells should be the number of antibodies in the tube (Video 3). Additionally, AF extraction should be tested to determine if marker resolution can be improved (Video 4).

The following protocol is divided into three sections: Discover, Distinguish and Designate.

Informed gating is a useful way to check panel quality. It is necessary to check that all populations of interest can be identified and to investigate how well they can be resolved (Fig. 10A, Video 6). The biology of the sample should be considered to ensure that populations seen in the sample are as expected and that markers are expressed on all the expected cell types. This protocol indicates whether the panel is ready for use on experimental samples. Additionally, dimensionality reduction algorithms can be useful tools for investigating panel quality, as they simplify the data for visualization while also exposing artefacts that may be missed through expert gating. Using t -distributed stochastic neighbor embedding (t -SNE; van der Maaten & Hinton, 2008; Fig. 10B), markers that are usually co-expressed should be checked to see they are found in similar regions of the t -SNE plot (Brummelman et al., 2019).

Background Information

The first step of building a successful large multicolor panel is good theoretical panel design. Full-spectrum flow cytometry panel design has been previously described (Ferrer-Font, Pellefigues, et al., 2020; Park et al., 2020). In summary, in the context of successful theoretical panel design, it is very important to have a clear experimental question and to know the biology of the markers that are included in the assay, including their expression and co-expression patterns. It is important to understand the instrument configuration (i.e., number of lasers on board) to know what the spectral signatures for each fluorophore will look like with the given configuration, as fluorophore brightness will vary depending on the excitation wavelength available. This information, coupled with the amount of spread (both given and received) of each fluorophore in the panel (detailed in Nguyen, Perfetto, Mahnke, Chattopadhyay, & Roederer, 2013), allows fluorophores to be optimally assigned to the different markers of the immunophenotyping panel.

After the panel has been theoretically designed, performance of QC steps is strongly recommended. This process is necessary to theoretically validate the panel before proceeding to a full experiment in order to reduce preventable issues as much as possible. It is advised to review the panel on a marker-by-marker and population-by-population basis, making sure that fluorophores inducing considerable amounts of spread (which could impair marker resolution) are allocated to non-co-expressed markers, and that dim markers receive a minimal amount of spread, while fluorophore brightness and antigen expression levels are well matched (high-expressing antigens with dim fluorophores and low-expressing antigens with bright fluorophores). To address potential issues, markers that are available with multiple fluorophores can be substituted to see if spillover spread can be minimized. Additionally, fluorophores that create (but do not receive) the most spillover can be designated to dump or viability channels. It is strongly recommended to complete panel optimization before working with an actual biological experiment, as it is a wise investment in terms of time, effort, and cost, without jeopardizing precious experimental samples.

Critical Parameters

Multiple factors influence the success of this protocol. The importance of good sample preparation—addressing sample complexity issues and the use of high-quality controls—must be emphasized.

Working with complex samples

When working with more complex samples than PBMCs (e.g., skin, tumor, fat tissue), the steps for panel optimization are the same, but there are some extra considerations.

Quality of single-cell suspension. An important consideration that is sometimes underestimated is the quality of the single-cell suspension that will be used to run the immunophenotyping panel. As different immune cell subsets have varying susceptibility to cell death, the single-cell suspension should have a viability of 80% or more to ensure proportional representation of the original sample (Costantini et al., 2003). Cell death can occur for different reasons, including how the sample has been treated before staining. Cryopreservation and harsh digestion protocols can affect sample quality and these procedures also need to be optimized. Indeed, when working with digested tissue, epitopes for the markers of interest should be verified to ensure the digestion protocol has not negatively impacted them. For example, if the same epitopes of interest exist in the spleen, one should compare marker staining and epitope preservation between digested and undigested spleen (Ferrer-Font, Mehta, et al., 2020).

Assessing instrument setup. Wherever possible, the instrument settings for fluorescence detector gains should remain unaltered from the Cytek Assay Settings (CAS). When using fluorescently labeled antibodies, this is achieved by carefully pairing expression levels to fluorophore brightness during panel design, and then by antibody titration to ensure fluorescence signals remain on-scale at the optimized settings. In certain applications where fluorescence intensity is not tunable, such as fluorescent reporter protein expression within the cells of interest, the instrument settings may need to be adjusted to accommodate off-scale fluorescence signal. Any such change will impact not only the spectral signature being accommodated, but also other signatures with emission in the same area. Reducing fluorescence gains may lead to increased similarity between spectral signatures, which can give rise to more spillover spreading error and thus negatively impact marker resolution (Ferrer-Font, Pellefigues, et al., 2020). See Support Protocol 2 for steps required to alter fluorescence gain settings to accommodate off-scale signals. If using something other than the Cytek Aurora, it will be necessary to optimize the settings based on manufacturer's recommendations.

Autofluorescence. Samples with heterogeneous (Fig. 7C) or very bright homogeneous AF signatures (Fig. 7B) can make unmixing challenging. To improve the accuracy of the unmixing and to improve marker resolution in the case of heterogeneous AF samples, multiple AF reference controls can be created for each of the different AF signatures present in the sample (as if they were individual fluorophores included in the panel). The steps required for this process are divided in three sections called Discover, Distinguish and Designate. The goal is to Discover unique spectral signatures within the unstained control through use of a raw N × N plots; Distinguish these unique signatures into separate SRCs by exporting each population as a new FCS file which can then be reimported; and Designate each signature as unique fluorophores within the software to be unmixed as if they were part of the original panel. Care must be taken to ensure only clearly unique signatures (with similarity index <0.98) with at least 300 events of similar fluorescence intensity are distinguished to ensure high quality SRCs are generated (for a detailed protocol, see Support Protocol 3 and Video 4). For very bright homogeneous AF, the steps outlined in Basic Protocol 3 and Video 4 can be followed to improve unmixing and markers resolution outcomes.

Controls

The quality of controls will directly translate to the quality of unmixing and the data obtained. It is therefore worth investing the time to optimize them. It is of utmost importance to include all necessary controls from the beginning of a project. A complete overview about controls can be found in Maecker & Trotter (2006). Below is a summary of the categories of controls that should be considered when optimizing a flow cytometry panel.

Unstained controls. The unstained control is meant for AF assessment and should have a clean signature with no contamination from other fluorophores and match the tissue or sample type being analyzed. If, for example, different tissue types are used in an experiment, multiple unstained sample controls must be used for each tissue type. Similarly, if samples are being treated differently (e.g., fixed/fresh or stimulated/unstimulated), an unstained control should be included for each condition. It is not advised to mix samples from different conditions and collect only one unstained control, as it may not be possible to have enough events to generate clear AF signatures.

Spectral reference controls. Appropriate single-stained SRC samples are required for optimal unmixing of the fully stained sample. The purpose is to provide a signature of each fluorophore to be used by the unmixing algorithm. Briefly, SRCs should have positive and negative populations that are clearly separated or a universal negative; positive populations should be brighter than the fully stained sample; the negative and positive populations should have identical AF characteristics; sufficient events for both populations should be collected; and the fluorescence spectrum of the positive control needs to be identical to the one in the fully stained sample. To fulfill these best practices, cells and beads should be compared and the best option should be used. Special considerations for viability staining are important, as live and dead cells have different AF signatures, which means the positive and negative controls will not have the same AF signature. To overcome this issue, it is possible to kill all cells in the viability SRC (e.g., heat-killing at 55°-70°C for 5-10 min), stain only half of them, and mix these with the unstained cells. In this case, all the cells will be dead and the AF will be the same for the positive and negative controls.

SS cell controls. SS cell controls are used not only as reference controls but also to assess the performance of each marker compared to the FS tube. SS cells are gold-standard for each antibody performance and are used to quantify the spread of the negative population and/or shifts in the positive signal and thus any subsequent loss of resolution of markers in the panel.

Gating controls. A fluorescence minus one (FMO) control is a sample stained with all fluorophores used in the experiment except one. Analyzing the FMO control for each fluorophore in the panel is not required for panel optimization, but FMOs can be used as a guide to set the boundary between positive and negative events if it is ambiguous. FMOs also aid in the assessment of spread between positive and negative events and are an important tool for assessing panel performance. A good alternative for large 20+ color spectral panels is the use of fluorescence minus multiple (FMM) controls, as recently described by Jensen & Wnek (2020).

Core versus FS panel

When optimizing a high-dimensional immunophenotyping panel, it is recommended to first complete Basic Protocols 1-4 with a core panel. A core panel is limited to only the essential lineage markers required to identify the cell types of interest. This will reduce confounding factors when attempting to understand the source of errors. Once a core panel has been optimized, additional markers can be added (including readout functional markers, intracellular markers, etc.) and optimized with the knowledge that issues are not originating from the core panel.

Fixation/permeabilization buffers

Intracellular staining procedures also require special consideration. It is important to source the fixation/permeabilization reagent that is most appropriate for the markers being detected (e.g., transcription factors, cytokines, or intracellular proteins) while also considering their relative locations (cytosolic or nuclear). The type of fixation buffer can influence the staining of both intracellular and surface markers. The type of fixative can have an impact in many aspects of staining: damaging the epitopes, altering the fluor stability and resulting in a different optimal titer, altering the background fluorescence without altering the positive signal (leading to reduced resolution between positive and negative populations), and altering the fluorescence spectral signatures (causing spectral signature mismatches between FS samples and SRCs, resulting in unmixing issues). It is recommended that antibodies be titrated using the same fixation/permeabilization buffer conditions used for the final staining of FS samples and SRCs.

Troubleshooting

The term troubleshooting is used when one or more issues are found in the panel and the source of these issues needs to be identified and steps included to rectify them. In this regard, the exact steps provided for validating the panel can also be applied for troubleshooting. By following these clear steps, the user will gain a better understanding of the quality of the panel and identify issues prior to acquiring experimental samples.

In general, 80%-90% of unmixing issues that arise can be traced to suboptimal controls. Therefore, it is important to use controls that are well characterized, high quality, and appropriate for the experiment (i.e., matched to the conditions of the experimental sample). This is particularly vital if they are going to be reused.

The checklist provided in Figure 11 aims to help users follow clear steps to optimize and troubleshoot their panels. By following these steps, users should be able to discover issues if they exist and have a pathway and alternatives to address and resolve them. Additional troubleshooting for general issues that may arise during panel optimization can be found in Tables 2-5.

Understanding Results

The example panel optimized using this protocol and presented here aims to identify conventional T cell subsets in peripheral blood of healthy participants. The participants were infected with low-dose human hookworm as part of a longitudinal study. It is well established that during early parasitic infection there is an increase in T-helper Type 2 immune responses (Th2) in the T cell compartments due to the primary role these cells play in host responses to parasites (McSorley & Loukas, 2010). However, the wider effects on the human immune system of long-term chronic parasitic infection with the gut-residing hookworm Necator americanus (Na) have not been investigated. Therefore, a panel was designed and optimized to look at the following conventional and unconventional T cell subsets of interest: CD4⁺ T cells (Th1, Th2, Th17, Th9, Th22, and Tregs), CD8⁺ T cells (Tc1 and Tc2), and innate-like T cells (γδ T, mucosal associate invariant T [MAIT], and natural killer T [NKT] cells). This panel can assess the frequency of these cell subsets as well as their memory and activation phenotypes, using both expert gating and high-dimensional data analysis.

Some characteristics of the fluorophore selection for a certain panel including the similarity and complexity indices of the panel (Fig. 12A) and the spectral signatures of fluorophores used (Fig. 12B) have been included. Although some fluorophores have a high similarity index, they have been allocated to different cell types and therefore the impact of the expected spreading error should be minimal (Fig. 12). A panel distribution table has also been included (Fig. 12C) to show the peak emission wavelengths for the fluorophores and markers assigned to the panel.

After following all the steps of Basic Protocols 1-4 for our panel shown here, and resolving any issues that arose (such as one tandem degradation or implementing the sequential staining for some of the markers), high-quality full-spectrum flow cytometry data as defined by clear resolution of all expected populations was achieved. The optimized staining protocol for this panel is detailed in the Supplementary Material. Sequential staining was applied, following the order given in the protocol, for markers that showed a reduction in positive signal in the FS compared to the SS control, starting at the marker with the greatest reduction and working toward the one with the least reduction. This approach provided satisfactory results and did not require the testing of other combinations. To reduce staining procedure time, further optimization would be required to determine whether some markers could be added at the same time instead of using all of them sequentially.

A clean and clear positive population can be detected for all markers in the panel, with no signal resolution loss when all antibodies are combined. In some cases, such as for the activation markers, the use of FMOs is necessary to assist in evaluating gates to determine positive staining with confidence. All populations of interest could be found using expert gating, and the populations resemble the expected expression patterns and frequencies. Additionally, high-dimensional data analysis algorithms were successfully used without the appearance of artefacts, confirming the high quality of the data.

Time Considerations

The time needed for designing, optimizing, and analyzing a high-dimensional flow cytometry panel can be highly variable and can depend on the assay complexity, number of markers, wait-time required for reagents (geographically dependent), sample access and frequency of sample delivery (particularly for patient samples), duration of the disease model being investigated, and more. As an example, it took five months to optimize the digestion, design, and optimization of a 23-color spectral flow cytometry panel in gut tissue (Ferret-Font, Mehta, et al., 2020), whereas one month may be sufficient for a PBMC panel where digestion does not need optimization. It is therefore advisable to develop certain core panels that can be applied to several experimental questions and models.

Acknowledgments

This work was enabled by the Hugh Green Cytometry Centre at the Malaghan Institute of Medical Research. We wish to thank the Hugh Green Foundation, the Marjorie Barclay Trust, the Dinies Foundation and the HRC Independent Research Organisation (18/1003) for funding this study. We are also very grateful to the NIH Core Tetramer Facility for supplying the tetramers used in this panel and to Olivia Burn and Kathryn Hally for feedback on the manuscript.

Author Contributions

Laura Ferrer-Font : Conceptualization; investigation; methodology; project administration; writing–original draft; Sam J. Small : Investigation; methodology; writing–original draft; Brittany Lewer : Investigation; writing–review & editing; Katherine R Pilkington : Resources; writing–review & editing; Laura K. Johnston : Writing–review & editing; Lily M. Park : Writing–review & editing; Joanne Lannigan : Methodology; writing–review & editing; Maria C. Jaimes : Conceptualization; methodology; resources; supervision; writing–review & editing; Kylie M. Price : Funding acquisition; project administration; supervision; writing–review & editing.

Conflicts of Interest

Kate Pilkington, Lily Park, Maria C. Jaimes are employees of Cytek Biosciences, Inc., the manufacturer of the Aurora full-spectrum flow cytometer used in these studies. Joanne Lannigan is a paid consultant for Cytek Biosciences, Inc.

Data Availability Statement

Data available on request due to privacy/ethical restrictions.

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