Monitoring Cell Proliferation by Dye Dilution: Considerations for Probe Selection

Abstract

In the third edition of this series, we described protocols for labeling cell populations with tracking dyes, and addressed issues to be considered when combining two different tracking dyes with other phenotypic and viability probes for the assessment of cytotoxic effector activity and regulatory T cell functions. We summarized key characteristics of and differences between general protein and membrane labeling dyes, discussed determination of optimal staining concentrations, and provided detailed labeling protocols for both dye types. Examples of the advantages of two color cell tracking were provided in the form of protocols for: (a) independent enumeration of viable effector and target cells in a direct cytotoxicity assay; and (b) an in vitro suppression assay for simultaneous proliferation monitoring of effector and regulatory T cells.

The number of commercially available fluorescent cell tracking dyes has expanded significantly since the last edition, with new suppliers and/or new spectral properties being added at least annually. In this fourth edition, we describe evaluations to be performed by the supplier and/or user when characterizing a new cell tracking dye and by the user when selecting one for use in multicolor proliferation monitoring. These include methods for:

1. Assessment of the dye’s spectral profile on the laboratory’s flow cytometer(s) to optimize compatibility with other employed fluorochromes and minimize compensation problems;

2. Evaluating the effect of labeling on cell growth rate;

3. Testing the fidelity with which dye dilution reports cell division;

4. Determining the maximum number of generations to be included when using dye dilution profiles to estimate fold population expansion or frequency of responder cells; and

5. Verifying that relevant cell functions (e.g., effector activity) remain unaltered by tracking dye labeling.

1. Introduction

The number of fluorescent dyes commercially available for cell tracking, and the subset useful for proliferation monitoring (reviewed in [1]), continues to expand rapidly, with new suppliers and/or new spectral properties continually being added [2–4]. Although diverse in their chemistries and fluorescence characteristics, these reagents can be grouped into two main classes based upon their mechanism of cell labeling. Dyes of one class, here referred to as “protein dyes”, react with proteins to form a covalent bond. Dyes of the other class, here referred to as “membrane dyes”, stably intercalate into the lipid bilayer of cell membranes via strong hydrophobic associations. The term “proliferation dye” will be used here to refer to dyes of either class that: (a) exhibit sufficiently good chemical and metabolic stability to partition approximately equally between daughter cells at mitosis; and (b) are sufficiently non-perturbing, even at high initial labeling intensities, to allow multiple rounds of cell division to be followed based on dye dilution.

Due to their stability of cell association, cell tracking dyes of both classes are often used for in vivo studies of cell trafficking and recruitment in contexts such as transplantation [5,6], infection [7,8], stem cell identification [9,10], and cancer immunotherapy [11]. Both dye types have also proven valuable for a wide range of in vitro studies including antigen presentation [12–14], mechanism and specificity of cytotoxic effector killing [15–17] (Subheading 3.6), and regulatory T cell activity [18,19]. Infectious agents [20,21], subcellular components (e.g., plasma membrane) [22,23] can also be tracked, as can the fate and bioactivity of cell-derived vesicles [24–26]. Combining fluorescent cell tracking dyes with stably expressed genetic markers has become increasingly common as the spectral choices available for probe types have increased. This strategy has been used to monitor extent and/or symmetry of cell division [27–29], something not possible with genetic markers alone, and also to detect active cycling in T cells responding to antigen, both before tracking dye dilution was evident and after daughter cells could no longer be distinguished from unlabeled cells [8].

Monitoring the proliferative status of stem/progenitor and immune cells is among the most common applications of both classes of cell tracking dyes [1,2,18,19,27,30,31], due to the significant limitations associated with alternative methods. Using dye dilution to assess extent of cell division avoids safety and regulatory issues associated with tritiated thymidine incorporation, which is ill-suited for single cell analysis of mixed populations, detects only cells actively synthesizing DNA during the pulse, and doesn’t allow isolation of daughter cells for follow-on analyses (e.g., immunophenotyping, gene expression, proteomics, or functional assays). Non-radioactive DNA precursors (bromodeoxyuridine, BrdU; ethynyldeoxyuridine, EdU) are compatible with single cell detection by flow cytometry using a variety of fluorochromes but also detect only cells actively synthesizing DNA during the pulse, and because detection requires fixation and permeabilization, viable daughter cells cannot be isolated for functional studies. Although BrdU and EdU also dilute out as cells proliferate, toxicity associated with high levels of incorporation [32] means that they typically cannot be used to monitor more than one or two rounds of cell division before labeled cells can no longer be distinguished from unlabeled ones.

The spectral capabilities of flow and imaging cytometers and the range of choices available for cell tracking dyes have both expanded dramatically since the third edition of this chapter, increasing our ability to design multiplex, high parameter studies to decipher complex biological systems [5,8,14,22,25]. This makes it even more essential to know the advantages -- and limitations -- of given tracking dye(s) in order to select probe(s) well-matched to the needs of a given application, particularly as new dyes become available. Key considerations for obtaining bright, homogeneous labeling with tracking dyes differ considerably from those for labeling with antibodies, and also for protein dyes vs. membrane dyes [2,19,33,34]. Subheadings 3.1 and 3.2 describe the protocols used to label cultured U937 cells and human peripheral blood mononuclear cells (PBMC) for the work presented here. Assessment of spectral compatibility with available instrumentation configuration(s) and other fluorochromes has become even more critical as choices have expanded. This is discussed in Subheading 3.3, using data collected for three protein dyes (CellTrace^™ Violet, CellTrace^™ CFSE, and CellTrace^™ Far Red) and three membrane dyes (PKH67, PKH26, and CellVue® Claret) on two different flow cytometers to illustrate the impact of laser configuration, laser power, and optical filter choices on stain index for each dye, and as a consequence, the degree of spectral overlap/compensation required in other spectral windows.

Before a new cell tracking dye is used for proliferation monitoring, it is important to verify that rate of dilution is linearly correlated with rate of cell growth in a system where unstained cells are also present and where growth can be independently measured. Subheading 3.4 illustrates two methods for doing this using continuously dividing cultured tumor cell lines. In systems where both responders and non-responders are present, mathematical modeling [1,34,35] is often used to quantify: (a) extent of population expansion in response to a stimulus (typically reported as ‘Proliferation Index’ or ‘Expansion Index’); and/or (b) proportion of the initial population able to proliferate in response to that stimulus (typically reported as ‘Precursor Frequency’ or ‘Percent Divided’). In such cases, it is also important to know the maximum number of daughter generations that can be followed before highly divided dye-positive cells begin to overlap with unstained cells. This is illustrated in Subheading 3.5 for lymphocyte cultures proliferating in response to stimulation with anti-CD3 and anti-CD28 antibodies. Finally, it is essential for each laboratory to verify for their particular cell type that the final labeling conditions chosen do not alter the proliferative behavior or functional potency of labeled cells relative to unlabeled controls (Subheadings 3.5 and 3.6, respectively).

2. Materials

2.1. Cell Isolation and Cell Culture

1. Complete Medium (CM): RPMI 1640 supplemented with 10% heat inactivated fetal bovine serum (FBS), 25 mM HEPES, 0.1 mM non-essential amino acids, 1 mM sodium pyruvate, 2 mM fresh glutamine, 50 μg/mL gentamicin sulfate, and 5 × 10⁻⁵ M β-mercaptoethanol.

2. 10% Formaldehyde, methanol free, ultra-pure. Dilute to 2% in PBS (pH 7.4) and store refrigerated.

3. Hanks’ Balanced Salt Solution (HBSS) without phenol red, magnesium, or calcium. Store at room temperature until opened, then at 4–8 °C.

4. Histopaque®-1077. Store at 4–8 °C and use at room temperature.

5. IL-2 (Aldesleukin Proleukin for injection, NDC 53905-991-01; Novartis, New York, NY). Dilute stock (2.2 × 10⁶ IU/mL) in sterile HBSS to 1 × 10⁵ IU/mL, aliquot, and store at −80 °C. Do not refreeze after thawing; store at 4–8° C and discard thawed product after 7 days.

6. Phosphate Buffered Saline (PBS): Prepare 10X stock containing 1.37 M NaCl, 27 mM KCl, 100 mM Na₂HPO₄, and 18 mM KH₂PO₄. Adjust to pH 7.4 with HCl if necessary. Sterilize by 0.2 μm filtration and store at room temperature. Prepare 1X working solution by dilution of one part with nine parts tissue culture grade water.

7. Human peripheral blood mononuclear cells (hPBMC). Isolate hPBMC from heparinized peripheral blood or from TRIMA filters [19] using the laboratory’s standard density gradient fractionation protocol, with the addition of a final low speed wash (300xg) to minimize platelet contamination (see Note 1).

8. K562 Cell Line (see Note 2). Kind gift of Dr. Myron S. Czuczman, Roswell Park Cancer Institute, Buffalo, New York; also available for purchase (American Type Culture Collection, Manassas, VA).

9. U937 Cell Line (see Note 2). Generously provided by Paul Guyre, Lebanon, NH; also available for purchase (American Type Culture Collection).

10. 24-well polystyrene plates are useful for plating quadruplicate samples for kinetic studies.

11. 96-well U-bottom polypropylene stripwell plate consisting of 1.1 mL polypropylene tubes in strips of 8, racked in plates and sterile.

2.2. Antibodies

1. 1.0 mg/mL anti-CD3 (clone OKT3) and 1.0 mg/mL anti-CD28 (clone 28.2). Azide free, unconjugated preparations (eBioscience, San Diego, CA).

2. CD45 allophycocyanin (APC, clone 2D1) and CD45 Brilliant Violet 510 (BV510, clone HI30) (BD Biosciences, San Jose, CA).

2.3. Flow Cytometry Reagents

1. FCM Buffer: 1X PBS (pH 7.2) supplemented with 1% BSA, 0.1% sodium azide, and 40 μg/mL tetrasodium ethylenediaminetetraacetic acid.

2. 4′,6-diamidino-2-phenylindole (DAPI). Reconstitute powdered solid to 5 mg/mL in deionized water and store at 4–8° C. Prepare a working stock by diluting to 5 μg/mL in deionized water. Add 5 μL of working stock to each 100 μL of cells (0.25 μg/mL final) and let stand on ice for 30 min prior to data acquisition.

3. 7-Aminoactinomycin D (7-AAD). Reconstitute powdered solid to 1 mg/mL in PBS and store at −20° C. Prepare a working stock by diluting thawed 1 mg/mL stock to 100 μg/mL in PBS and store at 4–8°C. Add 4 μL of working stock to each 100 μL of cells (4 μg/mL final) and let stand on ice for 30 min prior to data acquisition.

4. Rainbow 6-Peak Calibration Particles for Instrument setup (Spherotech, Lake Forest, IL). Use for establishing reference intensities in employed fluorescence detectors as described in Subheading 3.4.1.

5. AccuCount Particles for cell counting (Spherotech). Use for single platform cell enumeration as described in Subheading 3.4.1.

6. PKH26 Reference Microbeads (Sigma-Aldrich, St. Louis, MO). Use for single platform cell enumeration as described in Subheading 3.4.1.

2.4. Cell Tracking Dyes

1. CellTrace^™ Violet (CTV), CellTrace^™ CFSE (CFSE), and CellTrace^™ Far Red (CTFR) (ThermoFisher Scientific, Waltham, MA); CytoTrack^™ Yellow (CYY) (Bio-Rad, Hercules, CA). CFSE is also available from other suppliers. Reconstitute lyophilized aliquots for cell labeling according to the manufacturer’s recommended concentrations: 5 mM for CTV, 5 mM for CFSE (see Note 3), 1 mM for CTFR, and 500X for CYY. Labeling chemistries are similar for all four dyes: non-fluorescent precursor compounds freely diffuse across the plasma membrane into the cytoplasm, where their acetate substituents are cleaved by non-specific esterases. This results in trapping of the charged fluorescent product and random protein labeling via covalent bond formation between free amino substituents and the dye’s succinimidyl esters.

2. PKH67, PKH26, and CellVue^® Claret (CVC) fluorescent cell linker kits (Sigma-Aldrich). Kits contain 1 mM dye solutions in ethanol and cell labeling diluent for general cell membrane labeling (Diluent C). CVC is also available from another supplier (Molecular Targeting Technologies, Inc., West Chester, PA). Store tightly capped at room temperature to avoid evaporation of ethanol and associated increases in dye concentration. If any dye solids are visible, sonicate dye stocks to redissolve before use and verify that dye absorbance remains within the range specified on the Certificate of Analysis available for each kit. These dyes are incorporated into membranes based on hydrophobic forces that drive partitioning from the aqueous phase in which the dyes are highly insoluble, into cell membranes where they are stably retained due to strong non-covalent interactions between their long alkyl tails and those of membrane lipids.

2.5. Flow Cytometers and Data Analysis Software

For routine data acquisition, any flow cytometer capable of acquiring forward and side scatter, DAPI, BV421, BV510, FITC, PE, and APC would be appropriate. Data in this chapter were collected using five different flow cytometers. For all figures showing flow cytometric data, axis labels follow the convention of ref. [36].

1. LSR II (BD Biosciences). Fitted with 355 nm (100 mW), 405 nm (25 mW), 488 nm (20 mW), 561 nm (50 mW), and 640 nm (40 mW) lasers. From the 488 nm laser, FSC and SSC were measured using 488/10 nm bandpass (BP) filters. From the 355 nm laser, DAPI fluorescence was measured using a 450/50 nm BP filter. From the 405 nm laser, CTV fluorescence was measured using a 450/50 nm BP filter, and CD45-BV510 was measured using 525/50 nm BP filter. From the 488 nm laser, CFSE and PKH67 fluorescence were measured using a 530/30 nm BP filter, and 7-AAD fluorescence was measured using a 695/40 nm BP filter. From the 561 nm laser, PKH26 fluorescence was measured using a 582/15 nm BP filter. From the 640 nm laser, CTFR, CVC, and CD45-APC fluorescence were measured using a 660/20 nm BP filter.

2. LSR Fortessa (BD Biosciences). Fitted with 355 nm (60 mW), 405 nm (50 mW), 488 nm (50 mW), and 640 nm (40 mW) lasers. From the 488 nm laser, FSC and SSC were measured using 488/10 nm BP filters. From the 355 nm laser, DAPI fluorescence was measured using a 450/50 nm BP filter. From the 405 nm laser, CTV fluorescence was measured using a 450/50 nm BP filter, and CD45 BV510 fluorescence was measured using a 525/50 nm BP filter. From the 488 nm laser, CFSE and PKH67 fluorescence were measured using a 530/30 nm BP filter and PKH26 fluorescence was measured using a 575/26 nm BP filter. From the 640 nm laser, CTFR and CVC fluorescence were measured using a 670/14 nm BP filter.

3. MACSQuant Analyzer 10 (Miltenyi Biotec, San Diego, CA). Fitted with 405 nm (40 mW), 488 nm (30 mW), and 635 nm (21.5 mW) lasers. From the 488 nm laser, FSC and SSC were measured using 488/10 nm BP filters. From the 405 nm laser, CTV fluorescence was measured using a 450/50 nm BP filter. From the 488 nm laser, CFSE and PKH67 fluorescence were measured using a 525/50 nm BP filter, and PKH26 and CYY fluorescence were measured using a 585/40 nm BP filter. From the 640 nm laser, CTFR and CVC fluorescence were measured using a 655–730 nm spectral window.

4. MACSQuant VYB (Miltenyi Biotec). Fitted with 405 nm (40 mW), 488 nm (50 mW), and 561 nm (100 mW) lasers. From the 561 nm laser, FSC and SSC were measured using 561/10 nm BP filters. From the 405 nm laser, CTV fluorescence was measured using a 450/50 nm BP filter. From the 488 nm laser, CFSE and PKH67 fluorescence were measured using a 525/50 nm BP filter. From the 561 nm laser, PKH26 was measured using a 585/15 nm BP filter, and CTFR and CVC fluorescence were measured using a 661/20 nm BP filter.

5. NovoCyte 3000 (ACEA Biosciences, San Diego, CA). Fitted with 405 nm (50 mW), 488 nm (60 mW), and 640 nm (40 mW) lasers. From the 488 nm laser, FSC and SSC were measured using 488/10 nm BP filters. From the 405 nm laser, CTV fluorescence was measured using a 445/45 nm BP filter. From the 488 nm laser, CFSE and PKH67 fluorescence were measured using a 530/30 nm BP filter, and PKH26 fluorescence was measured using a 572/28 nm BP filter. From the 640 nm laser, CTFR and CVC fluorescence were measured using a 675/30 nm BP filter.

6. FACS DiVa^™ 8.0.1 (BD Biosciences).

7. FCS Express 6.0 (De Novo Software, Glendale, CA).

8. FlowJo^™ v10.2 (FlowJo, LLC, Ashland, OR).

9. WinList^™ v8.0 (current version is v9.0) and ModFit LT^™ v4.0 (Verity Software House, Topsham, ME).

3. Methods

Virtually any eukaryotic cell can be stained with either class of tracking dye after a single-cell suspension has been obtained (see Notes 4 and 5). The labeling conditions described below have been successfully used to stain human peripheral blood mononuclear cells (hPBMCs) and cultured cells used for the proliferation and cytotoxicity assays discussed here, but are likely to require modification for other cell types, assay systems, or dye combinations (see Notes 6 and 7). Although CTV, CFSE, and CTFR are used herein to represent a typical protein labeling dyes; and PKH67, PKH26, and CellVue® Claret to represent typical membrane labeling dyes, many other tracking dyes are available and the principles described here also apply to optimization of staining conditions and flow cytometer choice for use of those dyes.

3.1. Cell Line and hPBMC Labeling with Protein Dyes (CTV, CFSE, CTFR, or CYY)

The method for CellTrace labeling described here is a simplification of the protocol described by Quah and Parish [33]. The method for CYY labeling is the manufacturer’s recommended protocol, as adapted for the labeling of cultured U937 cells.

1. Prepare a stock solution in anhydrous DMSO by adding the recommended volume to the provided pre-weighed single use vial, vortexing, and visually inspecting the vial to ensure complete dissolution. The manufacturer’s recommended stock concentrations are: 5 mM for CTV, 5 mM for CFSE (see Note 3), 1 mM for CTFR, and 500X for CYY.

2. Wash cells to be labeled twice in serum-free PBS (or HBSS). After resuspension of the cell pellet from the first wash, remove an aliquot for cell counting (see Note 8). After final wash, resuspend cells in serum-free buffer (see Note 9) at a final concentration of 1 × 10⁷ cells/mL (range for hPBMC: 0.5–50 × 10⁶ cells/mL), using a tube that will hold at least six times the volume of the cell suspension. For CYY labeling, pellet cells and carefully aspirate supernatant.

3. Immediately prior to cell labeling, prepare working solutions of CellTrace dyes (50 μM for CTV and CFSE; 10 μM for CTFR) by making a 100-fold dilution of the DMSO stock solution from step 1 in PBS (see Note 10). Prepare a 2X working solution of CYY by making a 250-fold dilution of the DMSO stock solution from step 1 in PBS.

4. For a final staining concentration of 1 μM CellTrace dye, add appropriate amount of working dye solution per mL of cell suspension: 20 μL/mL of cells for CFSE or CTV; 100 μL/mL of cells for CTFR (e.g., to stain 2 mL of hPBMC at a final concentration of 1 × 10⁷ cells/mL and 1 μM CTV, add 40 μL of working dye solution; see Notes 7, 11, and 12). For a final staining concentration of 2X CYY, add resuspend cell pellet from Step 3 in 100 μL of working dye solution per 10⁶ cells (e.g., to stain 5×10⁶ U937 at a final concentration of 10⁷ cells/mL and 2X CYY, resuspend cell pellet from step 2 in 500 μL of 2X working solution prepared in step 3).

5. Immediately triturate or vortex tube briefly to disperse Protein Dye throughout cell suspension. Incubate at ambient temperature (~21°C) for 5 – 15 min, with occasional mixing either manually or on a rotator, protected from light (see Notes 13 and 14).

6. Stop the reaction by adding a 5X volume of CM or a 1X volume of FBS and mixing well (see Note 15). Centrifuge at 400xg for 5 min at ~21°C and discard the supernatant.

7. Wash the cells twice with 5–10 volumes of CM. After resuspension of the cell pellet from the first wash, remove an aliquot for cell counting. After the final wash, adjust cell concentration to the desired cell density for functional testing during the final resuspension in CM.

8. Assess recovery, viability, and fluorescence intensity profile of labeled cells immediately post-staining to determine whether to proceed with assay setup (ref. [19]; see Note 16).

9. At 24 h post-labeling, verify that labeled cells are well enough resolved from unstained cells for purposes of the assay to be performed and that Protein Dye fluorescence can be adequately compensated in spectral windows to be used for measurement of other probes (Subheading 3.3; see Note 17). If samples are to be fixed and analyzed in batch mode, verify that loss of intensity due to fixation does not compromise ability to distinguish desired number of daughter generations (see Note 18).

10. Verify that labeled cells are functionally equivalent to unlabeled cells (Subheading 3.6; see Note 19).

3.2. Cell Line and hPBMC Labeling with Membrane Dyes (PKH26, PKH67, or CVC)

The method described here is illustrated in detail in ref. [34].

1. Wash cells to be labeled twice in serum-free PBS or HBSS (see Note 9), using a conical polypropylene tube (see Note 20) sufficient to hold at least six times the final staining volume in step 5. After resuspension of the cell pellet from the first wash, remove an aliquot for cell counting (see Note 8) and determine the volume needed to prepare a 2X working cell suspension (step 4 below) at a concentration of 1 × 10⁸ cells/mL for hPBMCs (range = 2–100 × 10⁶ cells/mL), or 2 × 10⁷ cells/mL for U937 cells. For example, to stain a total of 5 × 10⁷ hPBMCs at a final concentration of 5 × 10⁷ cells/mL, the volume of 2X cell suspension would be 0.5 mL.

2. Following the second wash in step 1, aspirate the supernatant, taking care to minimize amount of buffer remaining (no more than 15–25 μL) while avoiding aspiration of cells from the pellet (see Note 21). Flick the tip of conical tube once or twice with a finger to disperse the cell pellet in the small amount of fluid remaining, but avoid significant aeration since this reduces cell viability.

3. Prepare 2X working dye solution of PKH67, PKH26, or CVC. To a second conical polypropylene tube (see Note 20), add the same volume of Diluent C staining vehicle (provided with each membrane dye kit) calculated in step 1 for preparation of the 2X cell suspension. Add the appropriate amount of 1 mM ethanolic dye stock to the Diluent C (e.g., for a 2X working dye solution to give a final dye concentration of 5 μM after admixture with 2X cells in step 5, add 5 μL of dye stock to 0.5 mL of Diluent C). Gently vortex tube to ensure complete dispersion of dye in diluent, avoiding loss of fluid to cap or as droplets on walls. Proceed with Steps 4 and 5 as rapidly as possible (see Notes 22 and 23).

4. Prepare a 2X cell suspension by adding the volume of Diluent C calculated in step 1 to the partially resuspended cell pellet from step 2. Triturate 3–4 times to ensure complete dispersion of the pellet and proceed immediately to step 5. Avoid excessive mixing, which reduces cell viability.

5. Rapidly admix the 2X cell suspension prepared in step 4 into the 2X working dye solution prepared in step 3, triturating 3–4 times immediately upon completion of addition in order to achieve as nearly instantaneous exposure of all cells to the same amount of dye as is possible (see Notes 24 and 25).

6. After 3 min, stop the labeling by adding a 5X volume of CM (containing 10% FBS) or a 1X volume of FBS or other cell-compatible protein and mixing well (i.e., if 1 mL of cells was combined with 1 mL of dye, then add 10 mL of CM or 2 mL of FBS; see Note 26). Centrifuge at 400xg for 5 min at ~21° C and discard the supernatant.

7. Wash the cells twice with 5–10 volumes of CM, transferring to a clean conical polypropylene tube after the first wash for maximum efficiency (see Note 27) and removing an aliquot for cell counting. After the final wash, count and resuspend the cells in CM at the final desired cell density for functional testing.

8. Assess recovery, viability, and fluorescence intensity profile of labeled cells immediately post-staining to determine whether to proceed with assay setup (see Note 16).

9. Verify that labeled but non-proliferating cells (e.g., unstimulated control) are well enough resolved from unstained cells for purposes of the assay to be performed and that membrane dye fluorescence can be adequately compensated in adjacent spectral windows used for measurement of other probes (Subheading 3.3; see Note 28).

10. Verify that labeled cells are functionally equivalent to unlabeled cells (Subheading 3.6; see Note 19).

3.3. Spectral Characterization

Cell tracking dyes are commonly combined with each other, with fluorescent antibodies, and/or with genetic markers to enable: (a) in vitro or in vivo tracking of phenotypically defined subsets within heterogeneous populations (e.g., [5,22]); or (b) identification and characterization of cell populations that do or do not proliferate in response to a given stimulus (e.g., [8,18,37]). The multiplicity of spectral detection options available on most digital flow cytometers means that when selecting fluorochrome combinations for such studies it is important to evaluate the impact of candidate cell tracking dye(s) on the ability to simultaneously detect other common fluorochromes. This must be done in the context of each cytometer’s optical configuration, taking into account relative signal intensities expected from the different probes. Due to the extremely bright staining typically obtained with all cell tracking dyes, spectral overlap into channels excited by the same laser and signal arising from cross-laser excitation must both be taken into account. Where cells of interest are in limited supply, this can conveniently be done using cultured cells as described below.

1. Harvest logarithmically-growing U937 cells and label with tracking dye(s) of interest as described in Subheading 3.1 or Subheading 3.2.

2. Configure the flow cytometer to acquire data from all fluorescence detection channels and adjust FSC and SSC detector voltages to ensure that cells of interest are resolved from electronic noise/small debris and large aggregates.

3. Use unlabeled cells to establish voltage settings that place their measured fluorescence distribution fully on-scale within the first decade (preferably >95% of events with intensity ≥0). Using the same instrument settings, confirm that each labeled cell population is fully on-scale in the primary detection channel for the tracking dye being evaluated (see Note 29).

4. Use data analysis software to generate single-parameter histograms for each detection channel as shown in Figs. 1 and 2. Histograms should be serially-gated on: bivariate plots of Time vs. SSC-A to eliminate any events associated with unstable flow rate; FSC-A vs. FSC-H to eliminate doublet events (see Note 30); and FSC-A vs. SSC-A to discriminate cellular events from small debris.

5. Generate color-coded instrument- and dye-specific table(s) of quantitative compensation values. Access the spectral crossover/compensation matrix data for each dye from the analyses performed in step 4, copy-and-paste it into spreadsheet software such as Microsoft Excel, and apply ‘Conditional Formatting’ to the data cells (Table 1).

6. When fluorescence intensity distributions are partially or fully off-scale in the primary channel when the unlabeled distribution is placed in the first decade (e.g., CTFR and CVC at 10 μM), accurate values for % overlap or compensation cannot be determined (see Note 31). In such cases, raw intensity data can be used to determine which detectors will be most affected by overlap from a given dye, including detectors affected by cross-laser excitation can be obtained (Table 2; see Notes 32 – 34). It does not matter which measure of central tendency is used (e.g., median, mean, geometric mean, etc.), provided that all fluorescence measurements employ the same metric.

7. Assess effect of cytometer optical configuration on Stain Index. For any given dye, compensation and cross-laser excitation issues in non-primary channels must be balanced against ability to detect the desired number of cell generations in the primary channel. Fig. 3 illustrates the use of Stain Index to compare the extent to which cells stained with PKH26, CTFR, and CVC can be resolved from unstained cells on 4 different cytometers (see Notes 35 and 36).

Fig. 1. Spectral Profiles of Selected Protein Dyes on BD LSR Fortessa Cytometer.

U937 cells were labeled as described in Subheading 3.1 at final concentrations of 1 × 10⁷ cells/mL and either 10 μM CFSE, 10 μM CTV, or 1.25 μM CTFR (see Note 31). All samples were analyzed 24 h post-labeling so that CFSE could be run at voltage settings that placed unlabeled U937 cells in the first decade for each fluorescence detector, as would typically be done for a dye dilution study. When evaluating compatibility of a given tracking dye with other fluors, both spectral overlap and cross-laser excitation characteristics of dye-labeled cells must be considered (see Note 32). Compensation settings required under these conditions are shown in Table 1A and compensation settings for the same samples run on a BD LSRII are shown in Table 1B (see Note 33).

Fig. 2. Spectral Profiles of Selected Membrane Dyes on BD LSR Fortessa Cytometer.

U937 cells were labeled as described in Subheading 3.2 at final concentrations of 1 × 10⁷ cells/mL and either 10 μM PKH67, 10 μM PKH26, or 10 μM CVC. Since membrane dyes exhibit minimal division-independent dye losses, all samples were analyzed immediately post-labeling at voltage settings that placed unlabeled U937 cells in the first decade for each fluorescence detector. Again, both spectral overlap and cross-laser excitation characteristics of dye-labeled cells must be evaluated for compatibility with other fluors (see Note 32). Compensation settings required for PKH67 and PKH26 under these conditions are shown in Table 1A and compensation settings for the same samples run on a BD LSRII are shown in Table 1B (see Note 33). Spectral overlap data for CVC, which was off-scale high in all detectors associated with the 640 nm laser, are shown in Table 2 (see Note 34).

Table 1.

A. LSR Fortessa: % Compensation Required as a Function of Dye and Concentration
Laser	Detector	CTV 10 μM	CFSE 10 μM	PKH67 10 μM	PKH26 10 μM	CTFR 5 μM	CTFR 2.5 μM	CTFR 1.25 μM
	B530/30	0.2	Primary	Primary	3.3	0.0	0.0	0.0
	B575/26	0.1	41.9	28.4	Primary	0.0	0.0	0.0
	B610/20	0.1	21.4	12.9	81.4	0.0	0.0	0.0
	B695/40	0.1	7.8	5.2	42.1	0.5	0.2	0.2
	B780/60	0.1	4.3	5.1	24.2	0.3	0.1	0.1
	R670/14	0.1	0.0	0.0	0.8	Primary	Primary	Primary
	R730/45	0.1	0.0	0.0	0.5	206.9	62.7	59.8
	R780/60	0.0	0.0	0.0	0.2	82.8	19.9	18.4
	V450/50	Primary	0.0	0.0	0.1	0.0	0.0	0.0
	V525/50	19.8	1.9	1.3	0.1	0.0	0.0	0.0
	V585/15	4.4	1.5	0.5	0.6	0.0	0.0	0.0
	V610/20	3.0	1.3	0.4	0.7	0.3	0.1	0.1
	V660/20	1.1	0.6	0.3	0.4	12.9	3.2	2.9
	V710/50	0.7	0.5	0.3	0.3	9.1	2.2	2.0
	V780/60	0.3	0.3	0.3	0.2	3.7	1.0	0.8
	U379/28	0.2	0.0	0.0	0.1	0.0	0.0	0.0
	U450/50	2.0	0.0	0.0	0.1	0.0	0.0	0.0
	U740/35	0.2	0.1	0.0	0.2	0.5	0.2	0.2

B. LSRII: % Compensation Required as a Function of Dye and Concentration
Laser	Detector	CTV 10 μM	CFSE 10 μM	PKH67 10 μM		CTFR 5 μM	CTFR 2.5 μM	CTFR 1.25 μM
	B530/30	1.1	Primary	Primary		0.0	0.1	0.1
	B695/40	0.5	5.6	4.4		0.3	0.2	0.3
	R660/20	0.7	0.0	0.0		Primary	Primary	Primary
	R730/45	0.5	0.0	0.0		142.6	166.8	102.4
	R780/60	0.3	0.0	0.0		44.1	41.8	41.4
	V450/50	Primary	0.0	0.0		0.0	0.0	0.0
	V525/50	97.1	1.8	1.2		0.0	0.0	0.1
	V610/20	19.3	1.2	0.4		0.3	0.3	0.3
	V660/20	9.1	0.7	0.3		10.7	10.1	10.0
	V710/50	4.1	0.4	0.2		5.2	4.9	4.8
	V780/60	2.4	0.3	0.3		2.7	2.5	2.5
	U450/50	12.0	0.0	0.0		0.0	0.0	0.0
	U530/30	6.3	0.3	0.0		0.0	0.0	0.0
	Y582/15	0.8	0.1	0.0		0.0	0.0	0.1
	Y610/20	0.7	0.1	0.0		1.1	1.0	1.1
	Y670/30	0.4	0.0	0.0		0.9	0.8	0.8
	Y710/50	0.4	0.0	0.0		8.6	7.9	7.8
	Y780/60	0.5	0.0	0.0		5.2	4.7	4.7
0–10%	10–20%	20–40%	40–60%	>60%	Primary

Table 2.

Geometric Mean Intensities in Non-Primary Channels

Fortessa Laser	Detector	CTFR 10 μM	CVC 10 μM	LSRII Laser	Detector	CTFR 10 μM	CVC 10 μM	PKH26 10 μM
	B530/30	4	6		B530/30	4	4	71
	B575/26	4	6		B695/40	31	212	527
	B610/20	4	5		R660/20	9870	Primary	8
	B695/40	46	350		R730/45	9994	9980	10
	B780/60	27	237		R780/60	5839	2409	4
	R670/14	Primary	Primary		V450/50	1	0	0
	R730/45	9990	9979		V525/50	3	3	3
	R780/60	6938	9977		V610/20	32	3	14
	V450/50	3	3		V660/20	1445	81	10
	V525/50	4	3		V710/50	704	213	7
	V585/15	4	3		V780/60	358	148	5
	V610/20	24	4		U450/50	2	3	3
	V660/20	1069	91		U530/30	2	3	3
	V710/50	748	312		Y582/15	4	10	4353
	V780/60	303	172		Y610/20	138	22	Primary
	U379/28	3	3		Y670/30	114	228	136
	U450/50	3	2		Y710/50	1144	5147	1403
	U740/35	46	207		Y780/60	681	4235	945
				1st decade	2nd decade	3rd decade	4th decade	Off-scale hi

Fig. 3. Effect of Cytometer Configuration and Dye Negative Population Placement on Stain Indices for Selected Tracking Dyes (see Note 35).

U937 cells were labeled as described in Subheading 3.1 (1.25–10 μM CTFR) or Subheading 3.2 (10 μM PKH26, 2–10 μM CVC) at a final concentration of 1×10⁷ cells/mL and analyzed on the indicated cytometer at the time point when a dye dilution analysis would be started (T0 = immediately post-labeling, PKH26 and CVC; T1 ~24 h post-labeling for CTFR). Wherever possible, detector voltages were set to place an unstained control sample fully on-scale in the first decade and stained cells were analyzed under the same conditions (“Autofl. On-scale”). For samples where dye-positive cells were partly or completely off-scale when unstained cells were fully on-scale in the first decade, detector voltage was reduced to place dye positive cells on-scale and unstained cells were visualized using a bi-exponential scale (“Dye+ On-scale”). Stain Index was calculated using Formula 1 of Note 35.

(Panel A) As expected, the Stain Index for PKH26 was higher using the 50 mW 561 nm laser on the LSR II (excitation efficiency @ 561 nm ~65% of maximum) than using the 50 mW 488 nm laser on the Fortessa (excitation efficiency @ 488 nm ~20% of maximum), Also as expected, the Stain Index for CTFR decreased on both instruments as dye concentration used for labeling was decreased from 10 μM to 1.25 μM. Note however, that Stain Index values obtained for cells labeled with 1.25 μM CTFR were substantially higher on both cytometers when detector voltages were set to place unstained cells fully on-scale (Autofl. On-scale) than when voltages were reduced to place cells stained with 10 μM fully on-scale (Dye+ On-scale)(see Note 36).

(Panel B) U937 cells labeled with PKH26 or CVC and an unstained control were analyzed on the indicated instruments during the Bowdoin 2016 Annual Course in Flow Cytometry. Again, the Stain Index for PKH26 was higher when this dye was excited using the 561 nm laser on the MACSQuant VYB than when the 488 nm laser on the NovoCyte was used, whereas CVC was more efficiently excited by the 640 nm laser on the NovoCyte than the 561 nm laser on the VYB.

3.4. Evaluating Effects on Cell Growth and Linearity of Dye Dilution

Cultured cell lines are also useful for testing whether there is a linear correlation between rate of cell growth and rate of dye dilution for a particular cell tracking dye. Such systems allow direct measurement of growth by cell counting, whereas indirect measures such as tritiated thymidine uptake must be used to estimate extent of proliferation in more complex systems (e.g., hPBMC; see Table S1 of ref. [1]). Before evaluating linearity of dilution, however, it is necessary to verify that the staining conditions used do not alter cell growth rate compared with that of unlabeled cells. Subheadings 3.4.1 and 3.4.2 describe methods for testing dye effects on growth rate using (a) parallel cultures of stained and unstained cells or (b) co-cultures of stained and unstained cells, respectively. Subheading 3.4.3 illustrates how the resulting data can be used to evaluate the linearity of dye dilution.

3.4.1. Relative Growth Rate of Stained vs. Unstained Cultures using Counting Beads (Non-volumetric Cytometer)

Beads from different vendors may be employed (e.g., as indicated in Subheading 2.3.5 – 2.3.6), so long as their fluorescence can be distinguished from that of labeled cells in at least one detection channel. Although it is not necessary to know the absolute concentration of beads/mL, a sufficient number of beads should be present in the initial culture to allow a statistically meaningful number (>100) to be counted when high density cultures are harvested and analyzed by flow cytometry at later time points.

1. Harvest logarithmically-growing U937 cells and label with tracking dye of interest as described in Subheading 3.1 or Subheading 3.2, leaving an aliquot of harvested cells unlabeled to serve as a dye-naïve control. Adjust labeled and unlabeled cell suspensions to 2.5 × 10⁵ cells/mL in CM.

2. Prepare counting beads for addition to cell cultures by removal of azide and detergents. From the stock solution of fluorescent counting beads, remove an aliquot sufficient to give a concentration of 2.5–10 × 10⁴ beads/mL after admixing with the cell suspension to be used for culture initiation. (For example, if a bead concentration of 10 × 10⁴/mL is desired after admixing with 50 mL of cell suspension, aliquot a volume of bead stock containing at least 5 × 10⁶ beads.) Centrifuge the bead suspension at 400xg for 5 min at ~21°C, discard the supernatant, and wash twice with 10 mL CM, discarding the supernatant after each centrifugation. After the last wash, re-suspend beads in 1 mL of CM.

3. To the cell suspension from step 1, add washed beads from step 2 at the volume needed to give a final bead concentration of 2.5–10 × 10⁴ beads/mL but do not exceed 1% of the volume of cell suspension. (For example, if 5 × 10⁶ beads were washed and resuspended in 1 mL CM, adding 200 μL of bead suspension to 40 mL of cell suspension gives a final bead concentration of 2.5 ×10⁴ beads/mL).

4. Mix cell and bead suspensions to homogeneity and separately plate 1 mL of each of suspension into replicate wells of a 24-well flat bottom polystyrene plate for each time point in the kinetic study (e.g., to evaluate 6 time points in quadruplicate, plate 24 wells). Incubate labeled and unlabeled cell cultures in a humidified 37°C incubator with 5% CO₂ for 7 days.

5. Harvest replicate samples of plated cell cultures for each time point into 12×75 mm round bottom tubes by triturating each well to homogeneity with a P1000 pipette fitted with a clean, sterile tip, and place tubes on ice. For the T0 samples, this should be performed immediately after plating the suspensions from step 4, in order to ascertain the starting cell-to-bead ratio.

6. After collecting the samples for Day 3 and Day 5, triturate all remaining wells in the plate as in step 5. Remove and discard 500 μL of mixed cell/bead suspension from each well, and replenish with 500 μL of pre-warmed, CO₂-equilibrated CM in order maintain the cultures in log phase growth. After addition of CM, collect a post-dilution sample to verify that cell-to-bead ratio has not changed (i.e., no more than 5% difference from pre-dilution ratio).

7. Add an appropriate viability dye (50 μL of a 5 μg/mL DAPI solution or 40 μL of a 100 μg/mL 7-AAD solution) to each mL of harvested cells and incubate on ice for 30 min prior to data acquisition. Do not wash or further manipulate harvested cell suspensions, in order to avoid selective losses of either cells or beads.

8. Acquire harvested samples using a flow cytometer, collecting forward and side scatter characteristics (pulse area, height, and width) as well as all relevant fluorescence detection channels (pulse area). Ensure that the acquisition threshold is configured to allow fluorescent beads and cells to be collected using the same instrument settings. Using the gating strategy summarized in Fig. 4A, set a ‘Stopping Gate’ on R4 and strive to collect 2500 fluorescent beads.

9. Establish the appropriate detector voltage for each dye’s primary detection channel, placing unlabeled controls fully on-scale in the first decade and confirm that labeled cells are fully on-scale. If >5% of labeled cells are off-scale high, reduce voltage as needed to bring them on-scale and re-acquire unlabeled controls.

10. After voltages have been satisfactorily configured for each dye’s detector, acquire fluorescent Rainbow 6-Peak Calibration Particles and record mean intensity values from all peaks that are well resolved and fully on-scale in a given detector. To ensure consistent fluorescence intensities and enable direct comparison of data that is collected on separate days, use the recorded intensities as target values when re-establishing detector voltages on subsequent days of the experiment.

11. Using appropriate software and the gating strategy illustrated in Fig. 4A, quantify the number of beads acquired during sample analysis. Use serially-gated bivariate plots of: Time vs. SSC-A to include only events associated with stable flow rate (R1); FSC-A vs. FSC-H to eliminate doublet events (R2); and FSC-A vs. SSC-A to discriminate bead events (R3) from cellular events. Gating on ‘R1&R2&R3’, create a bivariate plot for any two detectors where beads are expected to be brighter than cells (e.g., Y710/50-A and V780/60-A) and establish a “beads-only” region (R4). Use the combined ‘R1&R2&R3&R4’ gate to generate a bivariate beads-only plot of Time vs. FSC-A and establish a rectangular region (R5) to exclude doublet bead events and provide a singlet bead count.

12. Separately, quantify the number of viable singlet cells acquired during sample analysis (Fig. 4A). Use an ‘R1&R2’ gated plot of SSC-A vs DAPI U450/50-A (or 7-AAD B695/40-A for CTV) to identify dead cells and beads (R6), and ‘NOT R6&R1&R2’ gated plot of FSC-A vs. SSC-A to distinguish small debris from live U937 cells (R7) and a ‘NOT R6&R1&R2&R7’ gated histogram of dye fluorescence (e.g., CTFR R660/20-A) to provide a singlet cell count and monitor dye dilution over time (Fig. 4B).

13. Calculate cell-to-bead ratio (Cells/Bead) for each acquired sample by dividing number of events in the ‘NOT R6&R1&R2&R7’ gate (cells) by number of events in the ‘R1&R2&R3&R4&R5’ gate (beads). For the unstained control and each dye labeled population, plot Cells/Bead vs. Time (Fig. 4C), and use the slope of log(Cells/Bead) vs. time to calculate population doubling times.

14. It is important to note that this method can only be employed in cell systems where phagocytes (e.g., monocytes, macrophages, or dendritic cells) are not present, as they can internalize the counting beads as illustrated in Fig. 5, and render it impossible to obtain accurate ‘Cell/Bead’ ratios,. Since the counting beads are fluorescent, it is relatively simple to determine by microscopy or ImageStream cytometry (EMD Millipore, Billerica, MA) whether cells are internalizing beads.

Fig. 4. Relative Growth Rate Determination Using Counting Beads (Subheading 3.4.1).

U937 cells were labeled as in Subheading 3.1 at the concentrations of CTV, CFSE, or CTFR indicated in Panel C and a final concentration of 1×10⁷ cells/mL. After addition of 3 × 10⁴ beads/mL to each sample, parallel cultures of labeled and unlabeled cells were cultured for 7 days, with addition of fresh CM on Days 3 and 5 to maintain logarithmic growth. On each day, quadruplicate wells for each test article were separately triturated to homogeneity, harvested, stained with DAPI (CFSE, CTFR) or 7-AAD (CTV) for viability assessment, and acquired on the LSRII cytometer.

(Panel A) Data files were analyzed using the gating strategy described in Subheading 3.4.1, steps 11 and 13, with a Stopping Gate (R4) set to 2500 beads.

(Panel B) Histogram overlays for viable U937 cells (‘NOT R6&R1&R2&R7’) present in unstained (filled distribution) and Day 0 – Day 7 CTFR-labeled cultures, normalized to the volume of sample associated with a bead count of 250. Inset: Days 0 – 3 on an expanded scale.

(Panel C) Cell/Bead ratio increased over time at similar rates in all samples, indicating that growth rate was not altered by labeling with any of the three dyes at the concentrations tested. Calculated doubling times ranged from a low of 24.8 h (unstained cells) to 26.3 h (CFSE stained cells).

Fig. 5. The Presence of Phagocytes Invalidates Use of Counting Beads for Relative Growth Rate Determination.

When PKH26 counting beads (red events) are added to unstimulated cultures of unstained hPBMC (Panel A, T = 0 h), they are readily distinguished from mononuclear cells (blue events). After 24 h in culture (Panel A, T = 24 h), hPBMCs with the highest FSC-A (presumed to be monocytes) increase in side scatter and become PKH26 positive (green events). Analysis of the T = 24 sample on an ImageStream®^X Mark II imaging flow cytometer (EMD Millipore) confirmed that the red, FSC^lowSSC^highPKH26^high events were singlet PKH26 beads (Panel B) while the green, FSC^highSSC^highPKH26^high events were phagocytes that had engulfed multiple PKH26 beads. Phagocyte ingestion of beads caused lymphocyte to bead ratio (ratio of FSC^lowSSC^lowPKH26^low blue events to FSC^lowSSC^lowPKH26^high red+green events) to increase from 1.53 at T = 0 to 2.25 at T = 24 although there was no cell growth during this period.

3.4.2. Relative and Absolute Growth Rate Determination in Co-cultures (Volumetric Cytometer)

Cytometers with volumetric cell counting capability allow growth rate to be monitored in absolute units (cells/mL) as well as relative units (% labeled cells in co-cultures) without the addition of counting beads.

1. Harvest logarithmically-growing U937 cells and label with tracking dye of interest as described in Subheading 3.1 or Subheading 3.2.

2. For co-cultures, add 1 × 10⁵ stained cells and 1 × 10⁵ unstained cells to a sufficient amount of CM to give a final volume of 10 mL and place in a 25 cm² culture flask. For parallel cultures, add 2 × 10⁵ stained cells in a sufficient amount of CM to yield a final volume of 10 mL. To a control flask, add 2 × 10⁵ unstained cells in a sufficient amount of CM to yield a final volume of 10 mL. Incubate in a humidified 37°C incubator with 5% CO₂ for 5–7 days.

3. Immediately after cells are placed in culture flasks, and at approximately 24 h intervals thereafter, mix well by triturating and withdraw a 1.0 mL aliquot for flow cytometric analysis. On Day 3 after trituration, remove the entire volume of cell suspension and dispense 2.5 mL back into culture flask, reserving 1.0 mL for pre-dilution analysis. Add 7.5 mL fresh pre-warmed, CO₂-equilibrated CM to maintain logarithmic cell growth, mix well by triturating, and withdraw a second 1.0 mL (post-dilution) sample.

4. Use the laboratory’s standard reference material(s) to ensure that the flow cytometer is giving reproducible intensity values in all detectors to be used (as described in Subheading 3.4.1). Configure the flow cytometer to acquire data from all fluorescence detection channels and adjust FSC and SSC detector voltages to ensure that cells of interest are resolved from electronic noise/small debris and large aggregates.

5. Use unlabeled cells to establish voltage settings that place their measured fluorescence distribution fully on-scale within the first decade in the primary detection channel for each tracking dye being evaluated. Using the same instrument settings, confirm that the labeled cell populations are fully on-scale. If more than 5% of the labeled cells fall off-scale high, decrease detector voltage to bring the labeled cells fully on-scale, re-acquire the unlabeled cells at the new setting.

6. Use appropriate software to generate a single-parameter histogram for each tracking dye in its primary detection channel. At a minimum, histograms should be serially-gated on: bivariate plots of Time vs. SSC-A to eliminate any events associated with unstable flow rate; FSC-A vs. SSC-A to discriminate cellular events from small debris and large aggregates; and additional viability and/or doublet gates as needed.

7. For data analysis, restrict the evaluated population to viable U937 on a bivariate plot of FSC-A vs. SSC-A, circumscribe the viable cell population with an elliptical region (P1), and establish analysis regions for labeled and unlabeled populations on the P1-gated fluorescence intensity histograms (Fig. 6A).

8. Using the gated histograms, record the following for each time point at which labeled and unlabeled cell populations remain non-overlapping (see Notes 37 and 38): (a) labeled cells/mL; (b) unlabeled cells/mL; (c) dilution factor from step 3 [pre-dilution cell count divided by post-dilution cell count], if appropriate; and (d) % labeled cells.

9. Using Microsoft Excel or similar software, compare growth rates for labeled vs. unlabeled cells by plotting % labeled cells vs. Time (Fig. 6B) and cells/mL vs. Time (Fig. 6C) to identify concentration(s) that do not impair cell growth.

Fig. 6. Relative and Absolute Growth Rate Determination using Volumetric Counting.

U937 cells were separately stained with each of the indicated tracking dyes as in Subheadings 3.1 and 3.2 (final concentrations: 1 × 10⁷ cells/mL and 10 μM dye) and placed in 1:1 co-culture with unstained cells as in Subheading 3.4.2. An unstained control at the same total density was cultured in parallel. Aliquots withdrawn at each time point were acquired on a MACSQuant 10 Analyzer during the Bowdoin 2014 Annual Course in Flow Cytometry. Fluorescence histograms for each dye, gated on light scatter (P1, Panel A) to eliminate debris and aggregates, were used to determine absolute counts (cells/mL) and % stained cells at each time point. Green histograms denote co-cultures; pink histograms denote the unstained control.

(Panel A) Baseline separation between stained and unstained populations was maintained through Day 5 for CTFR co-cultures (upper histograms) and those for all other dyes except CYY (lower histograms). For CYY, two approximately equal populations were still distinguishable in the Day 1 co-culture (lower middle histogram) but the “unstained” population was ~7-fold brighter than the unstained control and was beginning to merge with the stained population, which had decreased 2.7-fold in intensity from Day 0 to Day 1. By Day 2, only a single population was evident in co-culture (lower right histogram). CYY was the only dye for which unstained cells exhibited such a rapid and extensive right shift, suggesting that the cause was dye-specific and not due to trogocytosis or other cell-type specific transfer mechanisms (see Note 38).

(Panel B) Stable values for % dye positive cells over time indicated that cells labeled with CTV, CFSE, PKH67, PKH26, CTFR, and CVC cells were growing at rates similar to unstained cells in co-cultures. In contrast, % CYY stained cells increased dramatically from Day 1 to Day 2 due to loss of resolution between stained and “unstained” cells (see Panel A).

(Panel C) Plots of absolute cell counts vs. time (Day 3 value = average of pre- and post-dilution counts) confirmed that both stained (“pos”) and unstained (“neg”) populations grew logarithmically during the 5 day co-culture period, with doubling times (indicated in parentheses) similar to those of the unstained control culture (see plot legend).

3.4.3. Assessing Linearity of Dye Dilution

Once labeling conditions that do not perturb cell growth rate have been established, absolute cell counts obtained from parallel cultures or co-cultures (as described in Subheadings 3.4.1 and 3.4.2) can be used to evaluate the correlation between cell growth rate and dye dilution rate.

1. For each dye and time point (T) after culture initiation (T0), calculate:

   1. Fold Growth = count (T)/count (T0)

   2. Fold Dye Dilution = median fluorescence intensity (T0)/median fluorescence intensity (T)

2. Use Microsoft Excel or similar software to plot Fold Growth vs. Fold Dye Dilution for each dye of interest (Fig. 7; see Note 39). A perfect two-fold decrease in dye intensity at each cell division would give a linear correlation with slope = 1.0.

Fig. 7. Correlation between Growth Rate and Dye Dilution Rate (Subheading 3.4.3; see Note 39).

U937 cells labeled as described in Fig. 6 and unstained controls were placed in separate parallel cultures (Subheading 3.4.2; 2013 data) or in 1:1 co-cultures (Subheading 3.4.2; 2014 and 2015 data) for 4–5 days. Aliquots withdrawn at each time point were acquired on 3 different MACSQuant 10 Analyzers during the 2013 – 2015 Annual Courses in Flow Cytometry and analyzed as described in Fig. 6.

(Panel A) All protein dyes tested exhibited much greater dye dilution between T0 and T1 (~20 h) than could be attributed to cell growth alone, reflecting the early, division-independent intensity loss characteristic of this dye class. CTV and CTFR showed less division-independent dye loss than CFSE, as indicated by their lesser T0 – T1 dilution despite similar growth rates for all three dyes (1.3–1.6-fold increase in cell counts). During the remainder of the culture period dye dilution was linear for all three dyes and more closely reflected cell growth rate, with slopes slightly greater than theoretical (range: 1.4–1.8).

(Panel B) All membrane dyes tested exhibited relatively linear dilution from throughout the entire culture period, with slopes similar to or slightly less than theoretical (range: 0.6–1.2).

3.5. Further Considerations for Cell Proliferation Monitoring

Although cultured cell systems are useful for establishing spectral compatibility with other fluorescent probes (Subheading 3.3; Figs. 1 and 2) and linearity of dye dilution for a given cell tracking dye (Subheading 3.4; Fig. 7), additional considerations arise when the biology of interest includes both responders and non-responders (e.g., immune cell populations responding differentially to a specific stimulus). In such cases, mathematical modeling is used to quantify the frequency of cells within the starting population that go on to respond to the stimulus and/or overall expansion within the responding subpopulation. Because no currently available tracking dyes of either type give baseline resolution between daughter generations, selection of an appropriate proliferation dye requires knowing the number of daughter generations that can be reliably discriminated from undivided non-responders using a given dye. This is determined by several factors: (a) the highest tolerated labeling concentration, which will dictate the initial cell labeling intensity; (b) extent of division-independent dye loss for a given dye (Fig. 7A); and (c) effect of stimulation on the autofluorescence of dye-naïve cells. In addition, in situations where a distinguishable non-responder peak is not present, it is important to know whether the location of stained but unstimulated controls from the same time point can be used as a surrogate to establish the expected location of non-responders in the stimulated sample. Assessment of these considerations is illustrated here using hPBMC stimulated with anti-CD3 and anti-CD28 to generate a polyclonal T cell response.

1. Generate a preparation of hPBMCs from peripheral blood using the laboratory’s standard density gradient fractionation protocol, with the addition of a final low speed wash (300xg) to minimize platelet contamination (see Note 1).

2. Enumerate harvested cells using the laboratory’s preferred methodology.

3. Label hPBMCs with desired tracking dye (Subheadings 3.1. and 3.2). Reserve a sufficient number of unlabeled hPBMCs to employ as autofluorescence assay controls; generally, an equal number of dye-labeled and unlabeled cells are required.

4. Adjust unlabeled and dye-labeled cells to 2 × 10⁵ cells/mL in CM and divide both labeled and unlabeled hPBMC into two equal volumes. To half of each cell suspension add azide-free anti-CD3 (clone OKT3) to a final concentration of 1 μg/mL plus anti-CD28 (clone 28.2) to a final concentration of 0.5 μg/mL (stimulated culture). Leave the other half unstimulated.

5. Separately dispense 0.5 mL (1 × 10⁵ total cells) of each test article into quadruplicate wells of a 96-well U-bottom polypropylene stripwell plate for each time point to be evaluated, and incubate in a humidified 37°C incubator with 5% CO₂ for 96 h (see Note 40). For example, to evaluate 5 time points in quadruplicate, establish 20 stimulated wells and 20 unstimulated wells for each dye-positive hPBMC test article; in parallel, establish 20 stimulated wells and 20 unstimulated wells for unlabeled (dye-negative) control cells.

6. At each time point, remove the appropriate dye-positive and dye-negative stripwell tubes from the plate that correspond to the indicated time point, and return the remainder to the humidified 37°C incubator with 5% CO₂. Separate individual tubes from each selected strip, lightly vortex to re-suspend cells and then transfer the stripwell tubes to the bottom of individually-labeled 12×75mm tubes, without washing or manipulating the samples.

7. Add an appropriate viability dye (25 μL of a 5 μg/mL DAPI solution or 20 μL of a 100 μg/mL 7-AAD solution) to each tube containing 0.5 mL of cultured cells and let stand on ice for 30 min prior to data acquisition.

8. Acquire data on the flow cytometer as described in Subheading 3.4.1, steps 9 and 10.

9. Use appropriate software (e.g., WinList, FCS Express, or FlowJo) and the gating strategy shown in Fig. 8A to analyze acquired data. Use serially-gated bivariate plots of: Time vs. SSC-A to include only events associated with stable flow rate (R1); FSC-A vs. FSC-H to select single cells (R2) and eliminate doublets; SSC-A vs. DAPI-A (or 7-AAD-A) to exclude dead cells (R3); FSC-A vs. SSC-A to define cellular events (R4); and the relevant fluorescence parameter(s) vs. SSC-A to generate fluorescence histograms for unlabeled or tracking dye labeled lymphocytes (see Note 41).

10. For every experimental condition assayed, generate overlay histograms of unlabeled and dye-labeled fluorescence distributions from each time point and stimulation condition, as illustrated in Figs. 8B and 8C. Identify the intensity at which highly divided cells can no longer be distinguished from the background fluorescence associated with unstained but stimulated cells. This will determine the maximum number of daughter generations that can accurately be modeled when estimating the fraction of the original population responding to a given stimulus and/or the extent of population expansion during the response (see Note 37).

11. Use appropriate peak modeling software (e.g., ModFit LT, FCS Express, or FlowJo) to analyze the viable, singlet, lymphocyte dye dilution histograms acquired at each time point from unstimulated (Fig. 8B) and stimulated (Fig. 8C) cultures and obtain the best fit to each according to the principles described in ref. [34] (see Note 37). For each unstimulated control sample, record the mean intensity of the highest intensity (parental) peak reported by the model (MFI, Parental). For each stimulated sample, record the mean intensity of the highest intensity (non-responder) peak reported by the model (MFI, Non-Responders). To determine whether dye dilution occurs at the same rate in non-proliferating cells under stimulated vs. unstimulated conditions, plot MFI, Non-responders vs. MFI, Parental as shown in Fig. 9. For such a plot, a slope of 1.0 indicates that there are no metabolic or other differences causing a systematic difference in rate of dye dilution in stimulated vs. unstimulated cultures.

Fig. 8. Effect of Dye Choice on Division-Independent Dye Dilution and Ability to Resolve Highly Divided Cells from Unlabeled Cells (Subheading 3.5).

hPBMC were separately stained with the indicated tracking dyes as described in Subheadings 3.1 and 3.2 (final cell concentration: 1 × 10⁷ cells/mL for CTV, CFSE, and CTFR; 5 × 10⁷ cells/mL for PKH26; final dye concentrations as shown) and cultured in quadruplicate without (Panel B) or with (Panel C) anti-CD3 + anti-CD28 stimulation. Data were acquired on an LSR Fortessa flow cytometer; one representative result is shown for each test condition.

(Panel A) Data files were collected using the gating strategy shown, with a Stopping Gate of 50,000 events in R4. Representative data and gating regions are shown for one of four replicate 96 hour cultures.

(Panel B) Overlays show Day 0–4 histograms for viable singlet lymphocytes from unstimulated cultures. Since lymphocytes do not proliferate under these conditions, T0 – T1 intensity decreases represent division-independent dye loss. As was seen for U937 cells (Fig. 7A), CTV and CTFR showed much less division-independent dye loss than CFSE.

(Panel C) Overlays show Day 0–4 histograms for viable singlet lymphocytes (‘R1&R2&R3&R4’ gate) in anti-CD3 + anti-CD28 stimulated cultures. As previously observed ([33]), stimulated but unstained lymphocytes exhibit measurably increasing autofluorescence over time, particularly in the V450/50 channel. As a result, even though CTV exhibits less division-independent early dye loss than CFSE, the ability to resolve unstained cells from stained but highly divided cells on Days 3 and 4 post-stimulation is actually slightly worse for cells labeled with 1 μM CTV than for those labeled with 1 μM CFSE. In situations where it is important to maximize resolution of unstained vs. highly divided cells but CTV concentration cannot be further increased without impacting cell function (see Subheading 3.6), use of a longer emitting tracking dye (e.g., PKH26, CTFR, CVC) may prove helpful.

Fig. 9. Effect of Stimulation on Rate of Dye Dilution in Non-Proliferating Lymphocytes (Subheading 3.5).

ModFit LT v4.0 was used to fit the quadruplicate histograms acquired each day for unstimulated (Fig. 8A) and stimulated (Fig. 8B) hPBMC cultures as previously described ([34]). The mean value for median fluorescence intensity (medFI) reported by the best fit model for unstimulated quadruplicates was plotted against mean value for medFI reported by the best fit model for non-proliferating cells (“non-responders”) in the stimulated quadruplicates. Error bars (± one standard deviation) are shown but in many cases are smaller than the plotted symbols. The T0 value for each dye is shown as a filled symbol with remaining time points shown as unfilled symbols. Despite the much greater initial intensity drop for CFSE, all four dyes studied gave slopes close to theoretical (range 1.0–1.1). This indicates that if there is not a distinct peak corresponding to the non-responders, the unstimulated control can provide a good estimate of its position in the stimulated histogram.

3.6. Evaluating the Effect of Tracking Dye Labeling on other Cellular Functions

In addition to proliferation monitoring, cell tracking dyes can be used for measuring cytotoxic effector cell activity by flow cytometry. This method does not require radioactivity and has the advantage of being able to measure killing at the single cell level even when targets and effectors cannot be distinguished on the basis of light scatter. The protocol described here uses killing of a cultured cell line (K562) by lymphokine activated killer (LAK) cells as a model system, but the principles and general procedures are applicable to virtually any effector-target combination.

3.6.1. Generation of Stained LAK Effector Cells

1. Prepare hPBMCs from peripheral blood using the laboratory’s standard density gradient fractionation protocol, with the addition of a final low speed wash (300xg) to minimize platelet contamination (see Note 1). Enumerate harvested cells using the laboratory’s preferred methodology and adjust cells to 1 × 10⁷ hPBMC/mL in HBSS.

2. Label hPBMC with desired tracking dye according to the procedures described in Subheadings 3.1 and 3.2 (see Note 42).

3. Assess recovery, viability, and fluorescence intensity profile of labeled cells immediately post-staining to determine whether to proceed with assay setup (see Note 43).

4. Resuspend labeled hPBMC in CM at 3 × 10⁶ cells/mL (typically 5–10 mL total volume) and incubate upright in a T25 flask with 1,000 IU/mL of IL-2 at 37° C for 4 days to generate LAK effector cells. Establish a parallel flask of unstained hPBMC for use as assay and instrument setup controls (Subheadings 3.6.3 and 3.6.4).

5. On Day 4, harvest LAK effector cells, triturating to disperse any cell clusters into a single cell suspension. Wash once with 50 mL CM, count and resuspend at 1 × 10⁷ cells/mL in CM.

3.6.2. Labeling K562 Target Cells

1. On Day 4 of the LAK induction period, harvest logarithmically growing K562 targets (see Note 44). Wash cells twice with 50 mL HBSS, count and adjust to 1 × 10⁷ cells/mL in HBSS.

2. Label K562 cells with desired cell tracking dye according to the procedures described in Subheadings 3.1 and 3.2.

3. Assess recovery, viability, and fluorescence intensity profile of labeled cells immediately post-staining to determine whether to proceed with assay setup (see Note 43).

4. Wash CFSE-labeled K562 targets twice in 15 mL CM. Count and adjust to 1 × 10⁵ cells/mL in CM.

3.6.3. Cytotoxicity Assay

1. In a 96-well round bottom polypropylene plate, make triplicate serial 1:2 dilutions of the LAK effectors as follows: Pipet 200 μL of the stained LAK cell suspension into the first well, and 100 μL of CM into each of 7 adjacent wells. Serially transfer 100 μL of LAK cells from the first well to the second, then from the second to the third, etc. ending with a transfer of 100 μL from well 7 to well 8 and removal of 100 μL of cell suspension from well 8.

2. Add 100 μL of stained K562 targets to each well, creating effector to target ratios of 100:1, 50:1, 25:1, 12.5:1, 6.2, 3.1, 1.6 and 0.8:1 (total volume per well: 200 μL).

3. Add 100 μL of targets and 100 μL of effectors to the target-only and effector-only wells respectively, followed by 100 μL of CM (see Table 3 for recommended assay and staining/instrument setup controls). Incubate the plate at 37° C for 4 h (see Note 45).

4. After the incubation period has elapsed, label test wells directly in the 96-well plate on ice for 30 min with a saturating amount of anti-CD45 antibody (see Note 46).

5. Transfer the contents of each well into individually labeled 12×75 mm round bottom tubes compatible with the laboratory’s flow cytometer. Wash each well with 200 μL of cold FCM buffer and transfer wash fluid to the appropriate tube.

6. Wash each sample once with 3 mL of cold FCM buffer and resuspend in 150 μL of FCM buffer.

7. Add 8 μL of 7-AAD (100 μg/mL stock) for CTV-labeled LAK cells; or 10 μL of DAPI (5 μg/mL stock) for CTFR-labeled or unlabeled LAK cells. Let stand for 30 min on ice so 7-AAD or DAPI can equilibrate before initiating acquisition of flow cytometric data (see Note 47).

Table 3.

Recommended Assay and Staining/Instrument Setup Controls for Dual Cell Tracking Cytotoxicity Assay (Subheading 3.6)

	Cells	Label	Treatment	Comments
Assay Controls1	LAK effectors only	CTV, CVC, or CTFR	Incubate with assay samples	Negative control: Used to calculate spontaneous LAK cell death
	K562 targets only	PKH67 or CFSE	Incubate with assay samples	Negative control: Used to calculate spontaneous K562 cell death
	K562 (heat killed) + LAK effectors	PKH67 or CFSE CTV, CVC, or CTFR	Incubate with assay samples	Positive control: only appropriate for assessment by 7-AAD or DAPI exclusion, not bead enumeration
	LAK cells	None	Same E:T ratios as test samples	Staining control: Used to verify that tracking dye labeled cells kill equivalently to unlabeled cells 5
Instrument Controls2	K562 cells	None	None	Select voltage for LAK- channel 6
	K562 cells	PKH67 or CFSE	None	Select voltage for K562 channel; set color compensation for all other channels; set negative region in LAK channel
	K562 cells	CD45 PacBlue or CD45-BV510	None	Set CD45 threshold or gate to include both targets and effectors (K562 MFI < LAK MFI)
	LAK cells	None 3	None	Select voltage for K562neg channel 7
	LAK cells	CTV, CVC, or CTFR	None	Select voltage for LAK channel; set color compensation in all other channels; set negative region in K562 channel
	LAK cells	CD45 PacBlue or CD45-BV510	None	Color compensation
	LAK cells	7-AAD or DAPI	Heat killed4	Color compensation

¹Negative and positive Assay Controls are included in the experimental plate with test samples, or set up in parallel with the experimental plate, to verify that the expected biological outcomes can be recognized using the chosen instrument conditions.

²Instrument Controls are used to establish instrument voltages and compensation settings.

³For unstained LAK cell controls, it will be necessary to set up a separate culture of unstained hPBMC with IL-2 at the same time as the PKH67 or CFSE stained hPBMCs.

⁴To heat kill, incubate at 56°C for 30 min. K562 cells could also be used but light scatter properties after heat killing differ substantially from those seen after LAK killing.

⁵Needed only to establish optimized staining conditions for tracking dye when assay is first being implemented in the laboratory; not required on a routine basis.

⁶Use of unstained LAK to select high voltage for the CTV or the “far red” (CVC or CTFR) detectors would place unstained K562 cells midscale due to their much greater autofluorescence. Therefore, unstained K562 cells were used instead to maximize dynamic range.

⁷Use of unstained K562 to select high voltage for the “green” (PKH67 or CFSE) detector would place unstained LAK cells off-scale low due to their much lower autofluorescence. Therefore unstained LAK cells were used instead to maximize dynamic range.

3.6.4. Flow Cytometric Acquisition and Analysis

1. Establish appropriate voltage settings using autofluorescence and single color controls from Table 3 (see Notes 43 and 48).

2. Using the single color controls from Table 1, adjust compensation settings according to your laboratory and/or instrument manufacturer’s standard procedures.

3. Acquire data on the flow cytometer and analyze data using the gating strategy described in Fig. 5 of ref. [19] and summarized in Steps 4–9 below (see Notes 46 and 49).

4. Evaluate all acquired events on a bivariate plot of SSC-A vs. CD45-A, using a rectangular region (R1) to restrict the subsequent analysis to CD45+ cellular events (and bead events, if added; see Note 49).

5. Gate R1 inclusive events to a bivariate plot of FSC-A vs. SSC-A, and use an irregular region (R2) to exclude any contaminating debris, but include all live and dead target and effector cells. If present, bead events should be separately circumscribed by a rectangular region (R3).

6. A reciprocal gating strategy is applied to assess target and effector cell numbers and viability. This strategy is used because substantial differences in autofluorescence exist between targets and effectors (see Table 3) which renders it difficult to establish instrument settings that provide complete resolution between K562 targets labeled with Tracking Dye 1 (TD1^pos) and LAK cells that are TD1^neg. Much better resolution is possible by using a second tracking dye to identify LAK cells (TD2^pos). Events satisfying the Boolean definition of R1&R2 (CD45^pos cells) are gated to a bivariate plot of TD2 fluorescence vs. Viability Dye fluorescence (see Note 47). Use a rectangular region (R4) to identify all target cells (TD2^neg). Then construct a gated (R1&R2&R4) bivariate plot of TD1 fluorescence vs. Viability Dye fluorescence to enumerate live (R5) vs. dead (R6) TD1^pos target cells.

7. Use a similar reciprocal strategy to quantify live vs. dead effector cells. First, gate CD45^pos cells (R1&R2) separately to a bivariate plot of TD1 fluorescence vs. Viability Dye fluorescence and use a rectangular region (R7) to identify TD1^neg effector cells. Then construct a gated (R1&R2&R7) bivariate plot of TD2 fluorescence vs. Viability Dye fluorescence to enumerate live (R8) vs. dead (R9) effector cells.

8. If counting beads are employed in the assay, enumerate them first on a gated (R3) bivariate plot of any two fluorescence parameters that yield good separation between broad-spectrum fluorescent beads and any cellular events that may have contaminated R3. Discriminate the counting beads from cellular events with a rectangular region (R10).

9. Construct a gated (R3&R10) bivariate plot of TIME vs. FSC-A, and employ a rectangular region (R11) to circumscribe all singlet beads in chronological continuity.

10. Calculate % cytotoxicity using the formula below (see Fig. 10 and Note 49).

    $$\%\mathit{Cytotoxicity}=\frac{(TD{1}^{+}\mathit{Viability}{\mathit{Dye}}^{+}\mathit{events})}{\mathit{Total}\mathit{number}ofTD{1}^{+}\mathit{events}}$$
    Similarly, % viable LAK effector cells is calculated as:

    $$\%\mathit{Viable}\mathit{Effectors}=\frac{(TD{2}^{+}\mathit{Viability}{\mathit{Dye}}^{-}\mathit{events})}{\mathit{Total}\mathit{number}ofTD{2}^{+}\mathit{events}}$$

11. An alternative method uses a calculation comparable to the approach used in a standard ⁵¹Cr release assay, using region R11 to enumerate singlet beads and region R5 to enumerate live K562 targets (see Note 49 and ref. [19] for details). The number of non-viable effectors in the assay can be calculated similarly; using regions R11 and R9 (see Note 50). This may be helpful in troubleshooting if target cell killing is lower than expected and/or in longer term assays in which some effector cell death is expected.

Fig. 10. LAK Cell Mediated Killing of K562 Targets is Unaffected by Staining.

LAK cells were independently labeled with CTV, CVC, or CTFR as described in Subheadings 3.1 and 3.2 and incubated with K562 cells labeled with PKH67 or CFSE at the indicated effector-to-target (E:T) ratios for 4 h at 37°C. Test samples and controls (Table 3) were analyzed using the gating strategy described in Fig. 5 of ref. [19] and steps 4–7 of Subheading 3.6.4. LAK cells used in Panels A and B were derived from the same donor several years apart. (Panel A: reproduced from ref. [19], with permission) LAK-induced cytotoxicity of PKH67-labeled K562 cells was assessed for each condition as percent of target cells that took up 7-AAD. As an internal control, percentage of dead LAK effectors (green lines) was assessed at each E:T ratio and verified to be acceptably low and relatively constant. To determine whether CVC staining affected LAK cytolytic potential, parallel studies were performed using CVC stained (solid lines) or unstained (dashes) LAK effectors. The data indicate that LAK cells kill K562 cells in a concentration-dependent manner, and that labeling with CVC did not affect their function. Representative data are shown from one of two replicate experiments analyzed on an LSRII cytometer; data points represent the mean ± 1 standard deviation of triplicate samples. Final staining concentrations used: LAK effector cells - 5 × 10⁷ cells/mL, 5 μM CellVue^® Claret; K562 targets – 1 × 10⁷ cells/mL, 10 μM PKH67. CD45+ effectors and targets were identified by staining with anti-CD45 Pac Blue (see Note 46).

(Panel B) LAK-induced cytotoxicity of CFSE-labeled K562 cells was assessed as a function of protein dye (CTV or CTFR) and staining concentration used (CTFR). For each condition, target cell viability was determined based on percent of target cells that took up DAPI. The data indicate that LAK cells kill K562 cells in a concentration-dependent manner, and that at the staining concentrations used neither CTV nor CTFR affected LAK cytolytic function. Final staining concentrations used: LAK effectors – 1 × 10⁷ cells/mL, dye concentrations as indicated on plot; K562 targets – 1 × 10⁷ cells/mL, 10 μM CFSE. CD45+ effectors and targets were identified by staining with anti-CD45 APC (when CTV was used to label LAKs) or anti-CD45 BV510 (for CTFR-labeled or unlabeled LAKs; see Note 46). Representative data are shown from one of two replicate experiments analyzed on an LSRII flow cytometer; data points represent the mean ± 1 standard deviation of triplicate samples.