Multiparametric Analysis of Apoptosis by Flow Cytometry
Flow cytometry is the most widely used technology for analyzing apoptosis. The multiparametric nature of flow cytometry allows several apoptotic characteristics to be combined in a single sample, making it a powerful tool for analyzing the complex progression of apoptotic death. This chapter provides guidelines for combining caspase detection, annexin V binding, DNA dye exclusion, and other single apoptotic assays into multiparametric assays.
This approach to analyzing apoptosis provides far more information than single parameter assays that provide only an ambiguous “percent apoptotic” result, given that multiple early, intermediate and late apoptotic stages can be visualized simultaneously. This multiparametric approach is also amenable to a variety of flow cytometric instrumentation, both old and new.
Key wordsApoptosis Caspase Flow cytometry Annexin V 7-Aminoactinomycin D Propidium iodide Pacific Blue Hoechst 33258
The importance of apoptosis in the regulation of cellular homeostasis has mandated the development of accurate assays capable of measuring this process. Apoptosis assays based on flow cytometry have proven particularly useful. They are rapid and quantitative; they provide an individual cell-based mode of analysis (rather than a bulk population) (1). The multiparametric nature of flow cytometry also allows the detection of more than one cell-death characteristic to be combined in a single assay. For example, apoptosis assays that utilize DNA dyes as plasma membrane permeability indicators (such as propidium iodide) can be combined with assays that assess different cellular responses associated with cell death, including mitochondrial membrane potential and annexin V binding to “flipped” phosphatidylserine (PS) (2, 3, 4, 5). Combining measurements for cell death into a single assay has a number of important advantages; it provides simultaneous multiple confirmation of apoptotic activity (important in a process that has proven highly pleiotrophic in phenotype). It also provides a much more comprehensive and multidimensional picture of the entire cell-death process.
Recognition of the pivotal role of caspases in the death process has led to the recent development of assays that can measure these important enzymes in situ. Caspase activation represents one of the earliest easily measurable markers of apoptosis (6). In most cases, caspase activation precedes degradation in cell permeability, DNA fragmentation, cytoskeletal collapse, and PS “flipping”; caspases are in fact both signaling agents and mediators of these downstream manifestations of cell death. Combining fluorogenic assays of caspase activation with fluorescence-based assays for later characteristics of cell death (such as PS “flipping” and loss of membrane integrity) can provide a very information-rich view of cell death. It can be particularly helpful in distinguishing the “early” stages of cell death from later events, allowing better signal transduction studies in cells prior to the complete collapse of the cell structure (7, 8, 9, 10, 11).
Several fluorogenic assays for caspase activity have also been described, including the OncoImmunin PhiPhiLux system, the FLICA substrates, and the NucView substrates (12, 13, 14, 15, 16, 17). All of these assays have both advantages and drawbacks. In this chapter, we describe the combination of the PhiPhiLux caspase substrate system with two simultaneous assays for later stages of cell death, annexin V binding to “flipped” PS residues, and cell membrane integrity using a DNA binding dye (17). The PhiPhiLux caspase substrates have several characteristics that make them useful for integration with other “live” cell apoptosis assays; they are cell-permeable and possess good caspase specificity. They are also relatively non-fluorescent in the intact state and become fluorescent upon caspase cleavage, with a signal-to-noise ratio of roughly 40 between the two states. They are also based on fluorescent probes with spectral characteristics similar to commonly used probes like fluorescein and rhodamine; this makes them easy to combine with other fluorescent probes (15, 16, 17). The ability to observe and measure multiple apoptotic phenotypes in a single assay gives a powerful picture of the overall apoptotic process. It is applicable to both suspension cells by traditional flow cytometry, and adherent cells using laser scanning cytometry (17). This assay can take advantage of newer flow cytometers with multiple lasers, but is also accessible to older cytometers with a single 488 nm laser.
- 1.PhiPhiLux G1D2 fluorogenic caspase 3/7 substrate (OncoImmunin, Inc., Gaithersburg, MD): OncoImmunin manufactures a series of fluorogenic enzyme substrates that fluoresce upon cleavage of an incorporated consensus domain. The fluorogenic caspase 3/7 substrate (PhiPhiLux G1D2) consists of an 18-amino acid peptide corresponding to the recognition/cleavage sequence from PARP, a target for caspase 3/7 (18). The substrate is homodoubly labeled with one of several fluorophores (in this case, a fluorescein-like molecule) on opposite sides of the molecule; in this conformation, the fluorochrome molecules are in close physical proximity and the fluorescence of the resulting complex is largely quenched (16, 19). After the substrate enters a cell by passive diffusion and is cleaved by caspase 3 or 7, the unquenched fluorescent fragments will be largely retained on the side of the membrane where the cleavage took place (16, 19).
- (a)The PhiPhiLux nomenclature indicates both its substrate specificity and the conjugated fluorochrome. The first letter refers to the substrate specificity: G refers to caspase 3/7, E to caspase 1, L to caspase 8, J to caspase 6, etc. The first number refers to the conjugated fluorochrome: 1 is the fluorescein-like fluorochrome, 2 to the rhodamine-like molecule and 6 to the sulforhodamine-like molecule. So G1D2 is specific for caspase 3 with the fluorescein-like probe, and E2D2 is specific for caspase 1 with the rhodamine-like probe. R2D2 is a special case and refers to the Cy5-like molecule, with the caspase indicated beforehand (3-R2D2 for caspase 3). Excitation and emission spectra for all the fluorochrome conjugates (generically referred to as X1D2, X2D2, etc.) are shown in Fig. 1.
PhiPhiLux G1D2 spectrally resembles fluorescein and can be excited with the standard 488 nm argon-ion or solid state laser found on most flow cytometers. The excitation and emission spectra for this conjugate and others are shown in Fig. 1. PhiPhiLux G1D2 is spectrally compatible with propidium iodide or 7-aminoactinomycin D (which can be used for measuring apoptotic cell permeability) and either phycoerythrin- or allophycocyanin-conjugated annexin V (for the detection of PS “flipping” during apoptotic death).
- (c)The PhiPhiLux reagents are roughly 40-fold dimmer in the uncleaved state than following caspase activation. The expected signal-to-background ratio between unlabeled and substrate-loaded viable and apoptotic cells is shown in Fig. 2, where cycloheximide-treated EL-4 thymoma cells were labeled with PhiPhiLux G1D2 and analyzed by flow cytometry; the apoptotic cells possess one- to three-orders of magnitude higher fluorescence than the viable cells. Primary cell cultures may show somewhat lower levels of caspase activation than cell lines, with subsequent lower levels of substrate fluorescence; however, background fluorescence may be lower with these cells as well (see Note 1).
The PhiPhiLux reagents are also available with other fluorescent tags, including rhodamine- and sulforhodamine-like fluorochromes, and a proprietary Cy5-like fluorochrome that can be excited with a red laser. The excitation and emission spectra for these alternative fluorochrome conjugates are shown in Fig. 1. The rhodamine and sulforhodamine substrates can be readily excited using green or yellow lasers, including 532 and 561 nm sources. These lasers are becoming more widespread on commercial cytometers. See Note 2 for more information on these conjugates.
The PhiPhiLux reagents are commercially provided at concentrations of 5–10 μM in sealed aliquots and can be stored at 4°C prior to opening; once the ampule is opened, any remaining substrate should be stored at −20°C. Avoid repeated freezing and thawing. Shelf life at 4°C is approximately 3–6 months, over 1 year at −20°C.
Phycoerythrin (PE)- or allophycocyanin (APC)-conjugated annexin V (available from multiple sources, including Invitrogen Life Technologies, Carlsbad, CA): Annexin V can be conjugated to a variety of fluorochromes, and binds to apoptotic cells with “flipped” PS residues on their extracellular membrane leaflet. Damaged or necrotic cells with a high degree of membrane permeability can also bind annexin V to their intracellular membrane leaflet, despite their uncertain apoptotic nature; therefore, a DNA binding dye as a cell permeability indicator should always be incorporated into annexin V binding assays. Cells that are both annexin V and DNA binding dye positive may therefore be either advanced apoptotic or necrotic.
- 3.DNA binding dyes: Keep in mind that all of the DNA binding dyes described here have differing cell permeability characteristics. This will affect the ultimate data analysis (see Note 3).
Propidium iodide (PI) is an inexpensive and widely available intercalating DNA binding dye. PI excites at 488 nm and emits in the 570–630 nm range. Dissolve in deionized water at 1 mg/mL and store in the dark at 4°C for up to 3 months.
7-aminoactinomycin D (7-AAD) (available from Sigma Chemical Co., St. Louis, MO and Invitrogen Life Technologies) is an intercalating/groove binding DNA binding dye that also excites at 488 nm, but emits in the far-red, with an emission peak at approximately 670 nm. 7-AAD is a good alternative to PI where a longer wavelength probe is desired. 7-AAD is also somewhat more cell permeable than PI. Dissolve 7-AAD in EtOH at 1 mg/mL and store at −20°C. Solublized stocks are good for 6 months. Diluted stocks should be used within 24 h.
Red- and violet-excited DNA binding dyes: The proliferation of cytometers with multiple lasers has greatly expanded the fluorochromes available for apoptotic analysis. Several red- or violet-excited DNA binding dyes can be substituted for PI or 7-AAD to increase total fluorochrome capability or to reduce fluorescence compensation requirements. Hoechst 33258 is a widely available minor groove DNA binding dye that is well-excited by ultraviolet or violet lasers; it has cell permeability characteristics similar to PI. Prepare Hoechst 33258 as a 1 mg/mL stock in distilled water, and store at 4°C for up to 3 months. Sytox Red and Sytox Blue (both available from Invitrogen Life Technologies) are red- and violet-excited DNA binding dyes that can be used; they are somewhat more cell-permeable than PI. Both Sytox Red and Sytox Blue are sold as stock solutions at 5 mM in DMSO with storage at −20°C, and should be diluted immediately prior to use. Very cell permeable DNA dyes like Hoechst 33342 (distinct from Hoechst 33258), the DyeCycle dyes (Invitrogen Life Technologies), and DRAQ5 (Biostatus Limited, Shepshed, Leicestershire, UK) should probably be avoided for most cytometric analysis of apoptosis, since they do not discriminate viable from apoptotic cells clearly enough. However, they may allow recognition of apoptosis-associated chromatin damage by microscopy and scanning cytometry.
Complete medium: RPMI supplemented with 10% FBS, l-glutamine, and penicillin/streptomycin
Wash buffer: Dulbecco’s PBS (containing calcium and magnesium) supplemented with 2% fetal bovine serum. This is used for cell washing prior to DNA dye addition. The inclusion of divalent cations is critical for annexin V binding.
Flow cytometer equipped with one, two, or three lasers.
(Optional) GemStone analysis software (Verity Software House, Topsham, ME).
3.1 Combinations of Fluorochromes
This assay combines fluorescent labels for three characteristics of cell apoptosis, namely caspase activation, PS “flipping”, and cell permeability. There is considerable flexibility of fluorochrome selection for the investigator depending on the flow cytometric instrumentation available. Three possible combinations are described below, one for analysis on instruments equipped with a single 488 nm laser, a second for instruments equipped with dual 488 nm/red diode or red HeNe lasers, and a third for instruments equipped with a violet laser diode.
3.1.1 Single 488 nm Laser Instruments
PhiPhiLux G1D2 (similar to fluorescein): Detect this fluorochrome in the fluorescein or FITC detector on most commercial instruments.
PE-conjugated annexin V: Detect this fluorochome in the PE detector on most instruments. Apply fluorescence compensation to separate the PE signal from PhiPhiLux G1D2 and 7-AAD.
7-AAD: Detect this far-red emitting DNA binding dye in the far-red (or PE-Cy5) detector on most commercial instruments.
3.1.2 Dual 488 nm/Red Laser-Equipped Instruments
PhiPhiLux G1D2 (similar to fluorescein): Detect this fluorochrome in the fluorescein detector on most commercial instruments.
APC-conjugated annexin V: Excite this fluorochome with either a red diode or HeNe laser, and detect in the far-red range. Little fluorescence compensation is required to separate its signal from PhiPhiLux G1D2 or the DNA binding dyes described below, making post-acquisition analysis easier. Annexin V conjugates with Cy5 and Alexa Fluor 647 (which are spectrally similar to APC) can be analyzed in the same way.
PI or 7-AAD DNA binding dyes can be incorporated into a cell-death assay with PhiPhiLux G1D2 and APC-annexin V. Detect both in the far-red detector (usually with a mid-600 nm bandpass (BP) or longpass (LP) filter) on most flow cytometers.
Further substitutions: If a red-excited DNA dye like Sytox Red is used, move annexin V to another detector (such as the PE detector).
3.1.3 Triple 488 nm/Red Laser/Violet Laser Diode-Equipped Instruments
- 1.PhiPhiLux G1D2 (similar to fluorescein): Detect this caspase substrate in the fluorescein detector. Combine it with:
PI, 7-AAD or Sytox Red: Either a 488 nm or red-excited DNA binding dye can be used (multilaser cytometers are typically equipped with both red and violet laser sources).
Pacific Blue-annexin V: Pacific Blue is a relatively bright violet-excited fluorochrome, and is available in an annexin V conjugate. Pacific Blue does not overlap significantly into other fluorescent channels, and other fluorochromes do not overlap significantly into it, making it very applicable for multiparametric assays.
- 2.Another possible combination still uses the fluorescein detector for PhiPhiLux G1D2, but uses:
Hoechst 33258 or Sytox Blue: These DNA binding dyes use the violet laser for excitation. Sytox Blue is somewhat more cell-permeable than Hoechst 33258, which is roughly equivalent to PI.
APC-annexin V: A red laser can be used to excite APC-annexin V. This combination uses three lasers to excite three fluorochromes; as a result, virtually no spectral overlap occurs, and almost no fluorescence compensation is required.
3.2 Preparation of Cells
Harvest cell lines grown in suspension or cultured primary cells. Transfer cells to 12 × 75 mm cell culture tubes, and centrifuge at 400 × g for 5 min.
Decant supernatant. Maximum removal of the supernatant is critical; the volume of remaining supernatant should be as low as possible to cause minimal dilution of the caspase substrate. Although cells can be washed prior to labeling, performing the assay in the remaining complete medium supernatant will reduce the amount of incidental cell death occurring during the assay. If cells are obtained from clinical or other in vivo sources, they should be centrifuged and resuspended in a complete tissue culture medium (such as RPMI containing 10% FBS) prior to use, then centrifuged, and decanted as described above.
Label 0.5 to 1 × 106 cells per sample; increasing this number will saturate the detection reagents and reduce caspase and annexin V labeling efficiency. Adherent cells pose special challenges for apoptotic analysis due to the physical trauma and membrane damage that occur with cell dissociation; analysis in the adherent state by laser scanning cytometry is much preferable to flow cytometry under these circumstances (see Note 4).
3.3 Fluorogenic Caspase Substrate Labeling
Ensure that as much supernatant is removed, to maximize final substrate volume. Tap each tube to resuspend the cell pellet in the remaining supernatant. The supernatant in the tubes will be approximately 50 μL in volume (but not exceeding 100 μL).
Add 50 μL of the PhiPhiLux reagent to each tube and shake gently. The PhiPhiLux reagent should be diluted as little as possible for maximum detection, hence the need for minimal sample supernatant. PhiPhiLux reagent solutions are typically prepared at 10 μM; this will give a final concentration between 3 and 5 μM (in approximately 100–150 μL of total volume).
Incubate the tubes for 45 min at 37°C, in a water bath or an incubator. An incubator may be preferred if CO2 conditions are desired. For both optimal labeling and reasons of economy, the PhiPhiLux reagent can be titered and tested for use between 0.5 and 5 μM. However, this should be done with caution (see Note 5).
3.4 Annexin V Labeling
After 45 min of caspase substrate incubation, add the appropriate fluorochrome-conjugated annexin V (in this case, either PE or APC). Annexin V is generally available in suspension at concentrations ranging from 0.1 to 1 mg/mL. Cell labeling should be carried out at approximately 0.5–5 μg annexin V per sample. Therefore, add 5 μL of a 1 mg/mL annexin V solution to the tubes described in Subheading 3.3, step 3. Again, fluorochrome-conjugated annexin V labeling should be titered in advance of actual use.
Incubate at room temperature (or in a 37°C incubator if CO2 is desired) for 15 min.
Add 3 mL of wash buffer to each tube. Centrifuge at 400 × g for 5 min, and decant supernatant.
3.5 DNA Binding Dye Labeling
Prepare a solution of DNA binding dye in complete medium: PI at 2 μg/mL, 7-AAD at 5 μg/mL, Sytox Red or Sytox Blue at 5 μM, or Hoechst 33258 at 2 μg/mL.
Add 0.5 mL of the DNA binding solution to each of the tubes described in Subheading 3.4, step 3. Maintain samples at room temperature and analyze within 60 min (see Note 7).
3.6 Flow Cytometric Analysis
PhiPhiLux G1D2: This fluorescein-like caspase substrate is detected through the fluorescein detector on most flow cytometers (often with the designation “FL1”) using a 530/30 nm or similar narrow BP filter. The spectral properties of PhiPhiLux G1D2 is similar to fluorescein, requiring some spectral compensation when used simultaneously with PE or PI (and to a lesser extent with 7-AAD).
PE-conjugated annexin V: Like most PE-conjugated reagents, this reagent is detected through the PE detector on most flow cytometers (often with the designation “FL2”) using a 575/26 nm or similar BP filter. PE requires some spectral compensation when used with PhiPhiLux G1D2 and 7-AAD.
APC-conjugated annexin V: APC is excited with a red laser source and detected through the APC detector on many flow cytometers (sometimes with an “FL4” designation) using a 660/20 nm or similar BP filter. An advantage of APC in multicolor assays is its minimal need for color compensation; there is no significant spectral overlap between PhiPhiLux G1D2, PI, or 7-AAD. Cy5 or Alexa Fluor 647 conjugates are spectrally similar to APC, and can be analyzed in the same way.
Pacific Blue-conjugated annexin V: Pacific Blue is analyzed using a violet laser; most instruments so equipped have at least two detectors aligned to this laser source. A 450/50 nm or similar filter is typically used to detect this fluorescent probe. Cascade Blue and Alexa Fluor 405 are spectrally similar to Pacific Blue, and are analyzed in the same way.
PI: This DNA binding dye is very bright even at low concentrations, and has a broad emission range, requiring compensation when used with PhiPhiLux G1D2. It can be detected in either the PE (575/26 nm filter) or far-red detection channel (red 650 BP or LP filter). The second choice is preferable to reduce spillover into the fluorescein detector. PE and PI can be analyzed together on older single laser instruments using the traditional PE detector (“FL2” detector, 575/26 nm) for PE detection, and the longer PE-Cy5 detector (“FL3” detector, 650 LP dichroic, or 675/20 nm) for PI. However, the close proximity of their spectra makes this analysis difficult. Substitution of PI with 7-AAD is preferable. PI is highly charged, and will contaminate instrument tubing, causing unwanted “shedding” of the dye into later samples. After PI use, the instrument should be thoroughly cleaned with 10% bleach or similar detergent to remove the dye.
7-AAD: This DNA binding dye is not as bright as PI and emits in the far-red, allowing its detection in the far-red channel on most single laser flow cytometers (the PE-Cy5, or “FL3” detector) with a 675/20 nm BP, 650 LP dichroic or similar filter. Compensation will be required when used with PhiPhiLux G1D2 and PE.
Hoechst 33258: Hoechst 33258 is very bright, and can be excited using either an ultraviolet or violet laser source. It is detected through a 450/50 nm or similar filter. It will have minimal spectral overlap into other fluorochromes. Like PI, it is highly charged and will adhere tightly to instrument tubing; the instrument should be cleaned thoroughly with 10% bleach or other detergent after use.
Sytox Red and Sytox Blue: These dyes can be analyzed using the conditions for APC and Hoechst 33258, respectively. Both are very bright, and are somewhat more cell-permeable than PI or Hoechst 33258.
3.7 Gating for Flow Cytometry
Scatter gating: Many cell lines and some primary cells show a dramatic alteration in forward and side scatter measurements late in the onset of apoptosis. Forward and side scatter are approximate indicators of cell size and optical density, respectively, and reflect both the cell volume loss and intracellular breakdown occurring during apoptotic death. It therefore seems logical to draw a gate around both the scatter-viable population AND the scatter-shifted apoptotic cells, and look at caspase activation, annexin V binding and DNA dye uptake in this total population.
However, the scatter-apoptotic population is also usually at very advanced stage of apoptotic death; the cells are already positive for all markers. The advanced physical perturbation of the cells in this group can also produce positive, but highly variable labeling results, interfering with the identification of earlier apoptotic stages. It is therefore also useful to gate only on the scatter-viable cells, and examine early apoptotic markers such as caspase activation only within this group of cells. This dual approach allows an overall picture of both early and late apoptotic stages, as well as examination of the earliest apoptotic cells. It is therefore recommended that both gating approaches be applied to get a clear picture of the apoptotic process (Fig. 2).
- 2.Annexin V binding and DNA binding dye exclusion: Exclusion gating can also be useful for markers other than scatter. Annexin V binding and DNA dye uptake usually occur after caspase activation and are considered “later” markers of apoptosis. Therefore, subpopulations negative or positive for annexin V and DNA dye binding can be gated for discrimination of “early” and “late” apoptotic cells. The annexin V-negative DNA dye-negative cells can be gated as in step 1 to allow detailed examination of the earlier stages of apoptosis such as caspase activation (Figs. 3 and 4).
Differences in DNA dye permeability: DNA dyes are not completely interchangeable with regard to exclusion by apoptotic cells (see Note 3). For example, 7-AAD is somewhat more cell-permeable than PI and will label an earlier subset of apoptotic cells; the Sytox dyes will also label earlier apoptotic cells than either PI or Hoechst 33258. This will affect the overall analysis. For example, if 7-AAD-positive cells are excluded from the analysis (in an attempt to quantify very early apoptotic events), this dye’s greater cell permeability will result in a lower apparent number of caspase-positive cells that are DNA dye-negative than if PI were used instead. These differences should be kept in mind when analyzing these early apoptotic subsets.
Caspase substrate background fluorescence: Viable cells labeled with a caspase substrate will have somewhat higher background fluorescence levels than completely unlabeled cells. Care should be taken to identify both the viable and apoptotic fraction without using an unlabeled control as a cutoff.
3.8 Simultaneous Immunophenotyping
The protocol described in this chapter is very compatible with simultaneous antibody immunophenotyping of the “viable” subpopulations. For example, PE-conjugated antibodies against a marker of interest could be combined with PhiPhiLux G1D2, 7-AAD, and APC-annexin V labeling as a very stringent “filter” for the removal of dead cells from the phenotyping analysis. This is similar to the common inclusion of PI or another viability probe in cell phenotyping protocols, to exclude dead cells from the analysis; incorporating a multicolor apoptotic assay with immunolabeling for dead cell exclusion is even more powerful. While a natural extension of this method would appear to be the immunophenotyping of early apoptotic cells (such as caspase-positive/7-AAD-negative/annexin V-negative), this should be approached with great caution (see Note 8).
3.9 Sample Results
Forward and side scatter: Figure 2 shows a typical shift in forward and side scatter during apoptosis in EL-4 cells treated with actinomycin D. In this case, both the entire population (excluding debris) and the scatter-viable cells are gated, and subsequently analyzed for caspase activation, annexin V binding, and DNA dye permeability.
Fluorescence distribution of PhiPhiLux G1D2 labeling: Figure 2 also illustrates the typical signal-to-background ratio between “viable” and apoptotic EL-4 cells labeled with the PhiPhiLux G1D2 substrate (shown here without annexin V and DNA binding dye labeling). The caspase substrate was readily detectable in the fluorescein channel by flow cytometry, in this case on a BD FACSCalibur. The substrate is much less fluorescent in the uncleaved state; signal-to-noise ratios of 1- to 3-log orders of magnitude are normally seen between “viable” and apoptotic cells loaded with PhiPhiLux G1D2. Unlabeled cells are slightly less fluorescent than “viable” labeled cells; this background fluorescence can be more dramatic in some cells types and does not necessarily indicate caspase activity in viable cells.
It should be noted that the “viable” and apoptotic distribution based on scatter measurements does not strictly correlate with caspase activation. The scatter-viable cells have a large percentage of caspase-positive cells, indicating that cells activate caspases prior to gross changes in scatter morphology. In some cases, the scatter-apoptotic population may also have some caspase-negative cells. While some of these cells may be advanced apoptotic or necrotic cells with diminished or degraded caspase activity, there may also be viable cells in this population. Previous studies have shown that cells may undergo transient volume fluctuations very early in the apoptotic process, well before caspase activation. These results indicate the importance of gating on both the total scatter-viable/apoptotic population, as well as the scatter-viable only cells.
PhiPhiLux G1D2 and 7-AAD labeling: Figure 3 shows the addition of the DNA dye 7-AAD labeling to the PhiPhiLux G1D2 assay. The dot plots at the top of the figure show 7-AAD labeling versus caspase activation for both drug-treated EL-4 cells gated for either the entire population (left dot plot) or the scatter-viable cells. Even with only two probes for apoptosis, three distinct subpopulations were apparent: a “viable” population at lower left, a caspase-positive population that had not progressed to 7-AAD permeability (lower right), and a caspase-positive population that was permeable to 7-AAD (upper right). Sometimes, a fourth population is also apparent that is also labeled with 7-AAD, but had little caspase activity. If present, this fourth population of cells likely contained necrotic or advanced apoptotic cells, where caspases had leaked out of the cells, or were proteolytically digested. Another important potential source of this population is cells that have undergone apoptosis in the incubation period following PhiPhiLux labeling but prior to flow analysis. Cells in this region demonstrate the importance of analyzing cells promptly at the completion of the assay, since apoptosis is still occurring. It also illustrates the importance of minimizing cell trauma during the assay; centrifugations and pipet transfers should be kept to a minimum.
At this point, the investigator can either include in the analysis all cells based on scatter (left column), or only the scatter-viable cells (right column). Excluding the advanced apoptotics can allow better resolution of the early-stage apoptotic cells. In addition, DNA dye labeling can now be used to exclude the more advanced apoptotic cells for specific measurement of the earlier dying cells. The bottom row of histograms show caspase 3/7 levels in 7-AAD negative cells. Caspase activation clearly precedes DNA dye permeability in this cell type.
PhiPhiLux G1D2, 7-AAD and APC-annexin V labeling: Figure 4 shows the final simultaneous analysis of caspase, annexin V, and DNA dye in a single assay. The left dot plot shows the forward and side scatter profile for apoptotic EL-4 cells; the entire cell population is then gated into a dot plot for annexin V binding versus DNA dye permeability (middle dot plot). Either the entire cell population or the annexin V-negative 7-AAD-negative cells can then be displayed for caspase 3/7 activation. A significant population of caspase-positive cells is present even in the annexin V-negative 7-AAD-negative population; caspase activation again precedes both of these characteristics. Layering multiple apoptosis assays into a single multiparameter assay therefore allows a comprehensive assessment of the apoptotic process in a cell population.
- 5.Detection of multiple caspases by flow cytometry: The PhiPhiLux system can incorporate a number of both consensus peptides for different caspase specificities, and fluorochromes for flow cytometric detection. It is therefore possible to load cells with more than one PhiPhiLux reagent, if they possess specificity for different caspases, and if they can be spectrally distinguished from one another by flow cytometry. This is illustrated in Figs. 5 and 6, where three caspase substrates were loaded simultaneously into apoptotic EL-4 cells, along with the DNA binding dye Hoechst 33258. The three substrates used were modifications of the PhiPhiLux reagent described earlier. Cells were simultaneously loaded with PhiPhiLux L1D2 (specific for caspase 8, conjugated to a fluorescein-like fluorochrome), PhiPhiLux R2D2 (specific for caspase 3/7, conjugated to a Cy5-like probe), and PhiPhiLux E2D2 (specific for caspase 1, conjugated to a rhodamine-like probe). The substrate concentrations were increased to allow simultaneous loading with all three substrate conjugates while maintaining the 3–5 mM concentration specified in Subheading 3.3, step 2. A BD LSR II equipped with 488, 561, and 405 nm lasers was used to excite this combination of fluorochromes (Fig. 5). The 561 nm laser was used to excite the rhodamine-like substrate, and provided adequate excitation for the Cy5-like substrate as well. Cytometers equipped with 532 and 561 nm lasers are now commercially available and becoming more common, giving access to these alternative substrate conjugates (Fig. 1). The rhodamine caspase 1 substrate used in this example was readily excited at this wavelength (Fig. 5).
- 6.Multiple caspase results and probability state analysis: Figure 6 shows the three-caspase activation profile, gated for the apoptotic (Hoechst 33258-negative) cell population. Caspase activation was clearly not simultaneous; caspase 1 and 8 are activated first, followed by caspase 3. This was confirmed using the probability state analysis software GemStone, which plots changes in flow cytometric parameters as functions of time, relative to a computer model (see Chapter 2 by Bagwell for discussion on Probability State Modeling, this volume). This analysis is shown in Fig. 7, where caspase 1 and 8 are upregulated prior to caspase 3. By using multiple lasers and caspase substrates conjugated to multiple fluorochromes, multiparametric assays for apoptosis can become much more informative. This modification to the multiparametric cell-death assay allows an even earlier stage of cell death to be distinguished and identified. Rather than just assaying for cell viability, investigators can collect important information about the signal transduction and effector processes of apoptotic death.
These collective results are consistent with many immune cell types and established cell lines; however, wider variations in apoptotic phenotype between different cell types should be expected (see Note 9).
Controls: Good “viable” and apoptotic controls are important for apoptotic analysis of apoptosis, and should be used, especially when a new cell type or apoptotic stimulus is being investigated. Where possible, an untreated negative control and an independent positive control should be included, the latter being induced by an agent other than that under study (such as a cytotoxic drug). The EL-4 cells used in this study are a good example of a control system that is easy to maintain and is reliable. Samples with both the absence and presence of the PhiPhiLux reagents are also important to include as controls, since the substrate does possess some low, but detectable intrinsic fluorescence in the uncleaved state that can be erroneously interpreted as apoptosis without the appropriate control samples. However, unlabeled cells should not be used as a strict guide for gating on PhiPhiLux labeled cells; they only allow determination of the increase in background from PhiPhiLux labeling.
Fluorogenic caspase substrates with alternative fluorophores: Fluorogenic caspase substrates coupled to rhodamine-like, sulforhodamine-like, and Cy5-like fluorophores are also available. The spectra of these probes are shown in Fig. 1, as illustrated in Figs. 5 and 6. None of these probes excite well at 488 nm; the rhodamine and sulforhodamine probes require a green or yellow laser source (532 or 561 nm DPSS laser), and the Cy5 probes require yellow or red excitation. The green- and yellow-excited probes were originally designed for epifluorescence microscopes, which are usually equipped with mercury arc lamp filters that can provide 546 and 577 nm (green and yellow) excitation light. Many flow cytometers are now equipped with lasers in this range, however, making these probes potentially useful for cytometric analysis. Using the rhodamine and sulforhodamine probes limits DNA dye choices; the spectrum of propidium iodide coincides too well with them, and therefore cannot be used simultaneously. All of these longer wavelength probes may give better sensitivity than the fluorescein-like versions; cellular autofluorescence is significantly reduced with green to red excitation (compared to 488 nm blue-green), so overall signal-to-background signal is likely to increase.
DNA binding dyes: All DNA binding dyes do not have identical cell permeability characteristics. Some DNA dyes will gradually cross the plasma membranes of even viable cells, while others are better excluded. These differences can affect the results obtained from the assay. For example, 7-AAD is somewhat more cell permeable than PI, and will give a greater percentage of apoptotic cells when compared directly to PI. Similarly, Sytox Blue is more cell permeable than Hoechst 33258, and will also identify an earlier set of apoptotic cells. This difference should be kept in mind while designing cell-death assays, and may dictate the use of 7-AAD when this property is desired. Highly permeable DNA binding dyes such as Hoechst 33342, the DyeCycle dye series, and DRAQ5, will enter cells and label their chromatin regardless of their viability state. This may limit their usefulness as apoptotic reagents for flow cytometry. They have however been used to identify morphological changes in chromatin during apoptosis by microscopy and laser scanning cytometry.
Multiparametric analysis of apoptosis in adherent cells: Flow cytometric analysis of apoptosis in adherent cell lines poses special challenges, since the removal of cells from their growth substrate may itself induce apoptosis. In addition, cell removal methods (such as trypsinization) can trigger false apoptotic indicators, such as aberrant annexin V binding in the absence of true cell death. By far the best solution to this problem is to utilize a laser scanning cytometer (LSC) for the analysis of apoptosis in these cell types; this specialized flow cytometer can perform cytometric analysis of cells on a flat surface, allowing minimal disruption during cell preparation (20). Several apoptosis assays utilizing caspase substrates using laser scanning cytometry have been described (17, 21, 22). The cell-labeling protocol is similar to that for suspension cells as described in Subheading 3.3, using cells cultured on tissue culture microslides as described previously (17).
Caspase substrate specificity and background: While the PhiPhiLux substrates seem reasonably specific for their target caspases, no synthetic substrate is exclusively specific for any particular enzyme. This should be kept in mind for any assay involving specific proteolytic activity. In general, a considerable excess of substrate will encourage low levels of non-specific cleavage, increasing the non-caspase background of the assay. Titration of the substrate to the lowest concentration able to distinguish activity may be necessary when the specificity of the assay is in doubt.
Annexin V: Calcium and magnesium are critical for annexin V binding; even brief removal of divalent cations after the binding reaction will result in rapid dissociation from PS residues. The cells must therefore remain in a calcium/magnesium buffer at all stages up to analysis, including all wash buffers.
Incubation periods and sample storage: All incubation periods and conditions are critical parameters for this assay, as is prompt analysis of samples following the labeling procedure. Insufficient incubation time for the PhiPhiLux substrates will result in poor labeling; prolonged incubation periods will increase the level of non-specific substrate binding and cleavage, resulting in high background fluorescence and decreased signal-to-noise ratios. In addition, prolonged storage of cells following removal of the surrounding PhiPhiLux substrate will eventually result in leakage of the cleaved substrate from the cell, despite its reduced cell permeability in the cleaved state. Overly long annexin V incubation periods will also increase the amount of non-specific binding to cells, making discrimination of “viable” and apoptotic cells more difficult. Although PI (and to a lesser extent 7-AAD) are relatively impermeant to viable cells, prolonged incubation will cause uptake even in healthy cells. If laboratory conditions do not allow prompt analysis of sample, cell-death assays involving fixed cells (such as TUNEL assays or immunolabeling of active caspases) should be considered as alternatives.
Simultaneous immunophenotyping of “viable” and early apoptotic cells: The protocol described in this chapter is readily amenable to the incorporation of antibody immunophenotyping along with the cell-death markers, resulting in a very sophisticated “screening out” of dead cells for measurement of receptor expression in “viable cells”. A potentially exciting extension of this method would appear to be the phenotyping of early apoptotic cells, positive for caspase expression, but negative for later markers. This method should be approached with care; from a cellular standpoint, caspase activation is probably not an “early” event in cell death, and many alterations in the plasma membrane may have occurred by this timepoint, resulting in aberrant antibody binding to cells as is observed in later cell death. Any cell surface marker expression results obtained by such methodology should be therefore be interpreted with caution.
Pleiotrophy in apoptosis: Apoptosis is a highly variable process involving multiple biochemical pathways; therefore, there are no universal morphological or physiological characteristics that are common to apoptosis in all cell types. Cell death in different cell types (even in physiologically or morphologically similar ones) may present very different phenotypes, and may not necessarily be detectable by the same assays. Multiparametric assays for apoptosis are very amenable to the nature of apoptosis, since the investigator is not limited to one characteristic of cell death. Investigators should also be willing to try other apoptotic assays to fully characterize their particular system.
The authors wish to acknowledge Veena Kapoor and Nga Tu Voong of the National Cancer Institute for excellent technical assistance, and Dr. Z. Darzynkiewicz of the New York Medical College for helpful discussion. Bill Godfrey, Jolene Bradford, and Gayle Buller at Invitrogen Life Technologies (formerly Molecular Probes) provided valuable technical information regarding fluorescent probes. Parts of this work were supported by intramural research fund provided by the Center for Cancer Research, National Cancer Institute.
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