Introduction
This chapter will describe common methods for measuring dead cells in culture using a plate reader that can be applied to high-throughput screening. This is an introductory review focused on the most frequently used methods for measuring the number of dead cells using a plate reader. We will describe basic assay methods and factors to consider when choosing an assay as well as the advantages and disadvantages of different methods. A separate chapter will describe HTS assays to measure apoptosis.
For all assays using cultured cells as a model system, it is valuable to know how many live and dead cells are present during or after the end of the experiment. This becomes especially important when cells are incubated for a period of time adequate to enable growth and division and thus change the total number of cells present. Relating live and dead cell numbers by a normalization process improves the statistical robustness of the assay. However, the use of cell number as an internal control is often overlooked in many cell-based assays such as reporter assays to detect expression of stress response genes or testing for other stress related events.
A frequent use of cells in culture is for a commonly used cytotoxicity assay where cells are exposed to a test compound and after some period of incubation, a marker is measured to reflect the number of viable cells present compared to positive (toxin) and negative (vehicle) control treatments. In addition to estimating the number of live cells, it can be of great value to measure the number of dead cells that have accumulated over the course of the experiment and to be able to distinguish between cytotoxicity and cytostasis or growth arrest. In some cases, estimating the number of accumulated dead cells may be more sensitive than measuring a decrease in a marker used to estimate viable cell number.
The two commonly used methods of estimating dead cells take advantage of the loss of membrane integrity and the ability of indicator molecules to partition into a compartment not achievable if the cell membrane is intact. As illustrated in the diagram in , assays used to detect dead cells include measuring the leakage of a component (usually an enzyme marker) from the cytoplasm into the culture medium or the penetration of an otherwise non-permeable dye into cells with a compromised membrane. This chapter will describe options for both general approaches for measuring dead cells.
Illustrates scanning electron micrographs of isolated rat hepatocytes. The left image is meant to depict an intact live cell and the image on the right depicts a dead cell with a damaged membrane. The loss of membrane integrity enables leakage of dead (more...)
Fluorescent DNA Binding Dyes That Penetrate Dead Cells
There is a large number of nucleic acid binding dyes that can be used to stain cells for microscopy or flow cytometry but have limitations for use with assays using plate readers to detect signal. Many dyes have somewhat similar properties which make it difficult to choose the most appropriate probe for a particular purpose. A general description of many of the nucleic acid binding dyes can be found in the following link to The Molecular Probes Handbook found on the ThermoFisher Scientific website.
There are many fluorescent DNA binding dyes to select from which are generally considered to be non-permeable to viable cells and can be used for detection of the accumulation of dead cells in culture using a multi-well plate format; however, there are a variety of factors to consider when selecting the most appropriate dye for assay development. The most important and practical factors to consider when choosing a dye include: the emission wavelength, selectivity for staining DNA, cell permeability, solubility at the vendor-recommended concentration, detection sensitivity and cytotoxicity. Fluorogenic DNA dyes that readily pass through the intact cell membrane and stain the nucleus of live cells should be avoided for consideration for measuring dead cells.
Chiaraviglio and Kirby (4) have recently reported on the evaluation of non-permeable DNA-binding dye fluorescence as a real-time readout of eukaryotic cell toxicity in a high-throughput screening format. They include a useful table summarizing the properties for many nucleic acid binding dyes as well as demonstrating the ability to multiplex fluorescent dead cell assays with other cell-based methods.
An important factor to consider when choosing any fluorescent dye is the emission wavelength spectrum. Knowledge of the excitation and emission spectra and the extent of any spectral overlap can be used to predict compatibility of two fluorescent assays and to select an appropriate filter set to avoid overlap of emission of different fluorophores. For example, a green-emitting DNA binding dye would be a logical candidate to multiplex with an assay detecting a red fluorescent protein. A table listing filter sets used for many of the nucleic acid binding dyes is included in the Chiaraviglio and Kirby reference (4). The ability to multiplex more than one fluorescent or luminescent assay provides flexibility during assay design. Measuring the number of dead cells is often used as a multiplexed internal control for other cell-based screening assays. Sequential multiplexing by recording data from a fluorogenic assay prior to addition of a second luminogenic assay chemistry can expand the possibilities for multiplexing.
The DNA binding dyes can be considered to be environmental sensors, meaning they change fluorescent properties after binding to various molecules. The various nucleic acid binding dyes may exhibit between a 20- to 1000-fold increase in fluorescence upon binding to double stranded DNA. That fold-increase can contribute to the relative sensitivity of detection of the number of dead cells. In most experimental conditions using a growing population of cultured cells in vitro, the quantity of DNA is proportional to the total number of cells present; however, changing culture conditions to induce rapid cell growth, to starve cells of nutrients, or induce differentiation to result in multinucleation may have a greater influence to change the amount of nucleic acid present in cells. The ideal situation for quantitatively detecting dead cells is for the dye to selectively bind to double stranded DNA. If the dye binds to double stranded RNA which may change under stimulatory or stressful culture conditions, using dyes that also bind to RNA can lead to artifacts and misinterpretation of results.
Even slight adverse effects of DNA binding dyes can limit their usefulness for real-time assay protocols where the dye is exposed to cells for extended periods of time. Dyes that cause cytotoxicity upon long term exposure to cells may be the result of partial permeability. Membrane permeability may depend on the cell type, the overall health of the cells or whether the dyes are substrates for efflux pumps that result in expulsion from the cytoplasm even if the dye does enter the cell. Reagent toxicity is not a problem if the dye is going to be used to stain cells for an endpoint assay protocol; however, cytotoxicity is critical to consider if cells will be cultured in the presence of the dye for an extended period to perform a real-time assay.
The use of DNA binding dyes for long term real-time detection of the accumulation of dead cells must consider if there is any influence of the assay reagent on the health or responsiveness of the cell model system. For example, for some cell viability assays, it is well known that reagents to estimate viable cell number (e.g. MTT and resazurin) can be toxic to the population of cells, even during a few hours of exposure (5). Similar reagent cytotoxicity effects are known for the DNA binding dyes. shows the effects of three different DNA binding dyes continuously exposed to four different cell types for 72 hours before measuring cell viability using an ATP assay.
Effect of DNA binding dyes on cell viability. Four different cell types were treated with the vendor recommended concentration of DNA binding dye and cell viability assayed at various times up to three days using ATP content as the marker.
The data suggest similar nucleic acid binding dyes may have different effects on cell viability and those effects can be cell type specific. Understanding the mechanism and use of these DNA dyes is therefore important to determine the best probe for the desired assay design. Like all potential toxins, the cytotoxic effect of assay reagent components can be expected to depend on the concentration, the duration of exposure, and the susceptibility of individual cell types. Appropriate controls (vehicle only without dye) are recommended to validate each dye and cell type combination to determine if there is a cytotoxic effect of the assay reagent. It is suggested to use the vendor recommended concentration as a starting point and test a range of concentrations of dyes with each cell model system to confirm there is not an artefactual cytotoxic or cytostatic effect.
Commercial Availability
Several examples of DNA binding dyes classified as nonpermeable to live cells are commercially available.
Propidium iodide: 1 mg/mL in water; ThermoFisher Catalog No. P3566
Hoechst 33342: 10 mg/ml in water; ThermoFisher Catalog No. H3570
SYTOX Green Nucleic Acid Stain: 5 mM Solution in DMSO; ThermoFisher Catalog No. S7020
YOYO-1 Iodide (491/509) - 1 mM Solution in DMSO; ThermoFisher Catalog No. Y3601
TO-PRO-3 Iodide (642/661) - 1 mM Solution in DMSO; ThermoFisher Catalog No. T3605
DRAQ7 far-red fluorescent DNA dye; Biostatus Catalog No. DR70250
GelRed 10,000X in DMSO; Biotium Catalog No. 41002
CellTox Green Cytotoxicity Assay: 1000X in DMSO; Promega Corporation Catalog No. G8741
Protocols and Sample Data
The following example protocols are based on using the CellTox Green Cytotoxicity Assay which contains a non-permeable asymmetric cyanine dye which binds the minor groove to stain DNA of dead cells. The CellTox Green Dye is optimally excited at 512nm with a peak emission at 532nm. The filter recommendations for detection are 485 ± 20nmEx and 520 ± 20nmEm. For a complete detailed description of protocols including: reagent preparation, determination of linear range, detection sensitivity for individual cell types, and endpoint or homogeneous assay protocols, refer to Promega Technical Manual #375.
The components of the CellTox Green Cytotoxicity Assay kit are provided frozen and include:
The DNA binding dye is provided frozen as a 1000X reagent dissolved in DMSO. Thaw the tube in a 37°C water bath, vortex to mix contents, and centrifuge briefly to collect all the liquid in the bottom of the tube.
The general goal of the protocol is to create a 1:1000 dilution of the DNA binding dye in the sample of cultured cells to be measured. There are optional protocols to prepare a working dilution in a balanced salt solution (Assay Buffer) and achieve a final 1:1000 dilution of dye depending on whether you choose to run the assay in a real-time mode (record data periodically over 1-3 days in culture) or an endpoint mode (measure once at the end of the experiment).
General Protocol / Real-Time Assay
- 1.
Add 10 μl of CellTox Green DNA binding dye per 5 ml of cell suspension in culture medium prior to seeding cells in the assay plates. That approach achieves a 1:500 dilution of dye in culture medium and allows for addition of test compounds (or vehicle control) as a 2X preparation in an equal volume of culture medium.
- 2.
Mix the tube containing the suspension of cells by inversion or gently vortex to ensure dye homogeneity prior to seeding into assay plates.
- 3.
Seed cell suspension in opaque walled assay plates (to avoid fluorescent signal crosstalk) with desired volume and cell number.
- 4.
Add test compound treatment (and vehicle controls) to each well using a volume of medium equivalent to that used to seed the cells so the final concentration of dye is a 1:1000 dilution of the original component.
- 5.
Measure fluorescence intensity at 485–500nmEx / 520–530nmEm at any time from 0 to 72 hours.
- 6.
Return the plate to the cell culture incubator between reads.
The following example in illustrates results from a real-time assay showing an increase in fluorescence with both higher Terfenadine concentration and longer incubation time. The increase in fluorescence indicates the accumulation of dead cells over a three day incubation. Cells treated with 100 µM toxin are stained within a few hours and maintain most of the fluorescent intensity over the three day period.
HepG2 cells treated with various doses of Terfenadine. CellTox Green Dye was added at time of Terfenadine dosing. Fluorescence was measured every hour over a three day incubation in a plate reader with an environmental chamber to control temperature at (more...)
Optional Protocol / Endpoint Assay
Addition of DNA binding dye can be used as an endpoint assay at the end of an experiment to stain dead cells and estimate how many are present. To assist in mixing of reagent and to avoid pipetting extremely small volumes, it is convenient to create a 1:500 dilution of dye in Assay Buffer and add a 1:1 vol:vol ratio of diluted dye reagent to the sample of cells in culture medium. Simply add the dye to a plate containing samples of cells exposed to test compound (and appropriate “no cell” and “no treatment/vehicle only” controls), mix using a plate shaker, wait 15 min, and record fluorescence intensity at 485–500nmEx / 520–530nmEm.
Optional Protocol / Multiplex Assay
The DNA binding dye in CellTox Green is functionally inert to the viable cell population which enables subsequent measurement of other parameters using a variety of compatible assay chemistries. One limitation to consider during multiplex assay design is the total volume available in the sample well. If you are performing a multiplex assay that requires addition of other reagents, you can include the DNA dye in the suspension of cells before dispensing into the assay plate or you can add a smaller volume of a more concentrated DNA dye reagent as long as the final concentration in the assay well is 1:1000 of the original dye component. For any lytic endpoint assay that will result in the second (multiplexed) reagent lysing the cells, the fluorescence value corresponding to the number of dead cells must be recorded first. The following example in shows a multiplex combination of assays by first recording the fluorescence resulting from the DNA binding dye to indicate the number of dead cells, followed by subsequent addition of a luminogenic reagent to quantify ATP as a marker for the number of viable cells. The multiplex combination of orthogonal assays to detect complementary cell health markers can be used to confirm overall results.
Multiplexing allows complementary measures of cell health. CellTox Green reagent was applied to bortezomib-treated K562 cells after 48 hours of exposure. Bortezomib is a known proteasome inhibitor chosen as a model toxin. Fluorescence associated with (more...)
Multiplex Assay Protocol
- 1.
Perform DNA binding dye detection of dead cells using either real-time or endpoint assay format as described above.
- 2.
Add a 1:1 vol:vol ratio of CellTiter-Glo Reagent to the samples containing CellTox Green Dye and experimentally treated cells.
- 3.
Place the plate on an orbital shaker and shake at 500–700 rpm to facilitate cell lysis and ATP extraction from the cells.
- 4.
Measure luminescence after 5–10 minutes.
Optional Protocol to Estimate Total Number of Cells
The total number of live plus dead cells at the end of an experiment can be estimated by including a detergent (Lysis Solution) with the DNA binding dye reagent to lyse and stain all of the cells present.
- 1.
Prepare a dilution of reagent by adding 20 µl CellTox Green Dye, 1,000X and 40 µl Lysis Solution (9% w/v Triton X-100) to 10 ml of Assay Buffer to create a 1:500 dilution of dye and a 1:250 dilution of Lysis Solution.
- 2.
Mix the tube containing the combined reagent by inversion or gently vortex to ensure homogeneity of the dye and detergent prior to adding to assay plates.
- 3.
Add a 1:1 vol:vol ratio of the combined reagent to the sample of cells to be measured.
- 4.
Mix the plate on an orbital shaker (700–900 rpm) for 1 minute to ensure homogeneity.
- 5.
Incubate at room temperature for 15 minutes, shielded from ambient light.
- 6.
Measure fluorescence intensity at 485–500nmEx / 520–530nmEm.
Controls for DNA Binding Dye Experiments
Background control (without cells). To determine background fluorescence, record fluorescence values from samples containing the recommended concentration of DNA binding dye in complete culture medium without cells.
Negative control (untreated cells). To establish a negative control, measure fluorescence from samples containing cells in complete culture medium treated with the same amount of vehicle/buffer used to deliver test compounds and containing the same concentration of DNA binding dye used for test samples.
Positive control (total cell number). To determine a value for the positive control representing all of the cells in the sample, a detergent (Lysis Solution: a 1:250 dilution of 9% wt:vol Triton X-100) is used to lyse the cells and enable staining of the entire population. To prepare positive control samples, add 4 µl of Lysis Solution per 100 μl of cells in culture medium, mix the samples using a plate shaker as described above, allow the samples to stand at ambient temperature and record fluorescence. Note: The dynamic range of the plate-reading fluorometer should be validated to ensure it is capable of reading over the complete range of fluorescence for the chosen cell number and probe concentration.
Troubleshooting / DNA Binding Dye
The optimal concentration of DNA binding dye for staining may vary among cell types and culture conditions. It is recommended to test a range of concentrations to optimize detection sensitivity.
Use appropriate controls including wells containing complete culture medium and vehicle only (without cells) to determine the background fluorescence.
Culture medium containing phenol red can quench overall fluorescence. The endpoint assay protocol that uses the provided Assay Buffer can reduce quenching and increase the signal-to-background ratio. The Assay Buffer should not be used to deliver the CellTox Green Dye for the kinetic real-time mode using longer incubation times.
Confirm appropriate filter selection is in place in the plate reading fluorometer.
Use opaque-walled 96-, 384-, or 1536-well tissue culture plates compatible with fluorometer.
DNA intercalating compounds (doxorubicin, actinomycin D, daunorubicin, etc.) may compete for binding of the DNA dye, thus reducing staining and leading to underestimation of cytotoxicity.
Extended incubations (more than 24 hours) can cause evaporation of culture medium and may lead to edge effects. Using only the inner wells of assay plates may mitigate this problem.
Monitoring the viable, dead and total cell number over the course of an experiment, enables determination of cytostatic vs cytotoxic effects of test compounds. Vehicle-treated control cells may continue to proliferate over the course of an experiment.
A general precaution for all DNA binding dyes is to protect the reagent from light.
Animal-derived products such as Matrigel Matrix which is a tumor extract may contain DNA and result in high background levels.
Use of DNA binding dyes to test for cytotoxicity resulting from transfection experiments is not recommended. Residual DNA from transfection mixture may result in high background fluorescence.
Markers That Leak Out of the Cytoplasm of Dead Cells into Culture Medium
Introduction
The presence of dead cells that have lost membrane integrity can be detected by measuring markers that leak from the cytoplasm into the culture medium. The most common marker used for this type of assay is lactate dehydrogenase (6, 7). Lactate dehydrogenase (LDH) catalyzes the conversion of pyruvate to lactate and in the process, converts NAD+ to NADH. The reducing capacity of NADH can be used to reduce a variety of substrate molecules into products that are either colored, fluorescent, or luminogenic. illustrates the general scheme and assay chemistry used to detect LDH-release from the cytoplasm of dead cells. An excess amount of lactate and NAD+ as substrates are delivered in a reagent mixture to drive LDH to generate pyruvate and NADH. The reducing power of NADH is used to convert the substrate (resazurin) into the fluorogenic product (resorufin). Colorimetric versions of this assay chemistry have used a tetrazolium compound as the diaphorase substrate which is converted into an intensely colored formazan product that can be measured using a spectrophotometer. Similarly, a luminometric assay can use a “pro-luciferin” substrate which is converted into a luciferin product that is linked to a firefly luciferase reaction to generate a luminescent signal. The colorimetric version of the assay was developed decades ago and lacks detection sensitivity. In addition, because of buffer incompatibility with live cells, it requires removal of culture supernatant to a different container to perform the assay. The fluorescent assay protocol is homogeneous and more sensitive than the colorimetric version. Advances in formulating the reagent to be compatible with viable cells enabled a homogeneous fluorescent protocol to be developed. The luminogenic version of the assay is far more sensitive than the fluorogenic version that enables sampling of 2-5 µl of culture supernatant at various times which can be stored frozen for future analysis of the trends of LDH release over time.
A schematic representation of the fluorogenic LDH-release assay chemistry. LDH from dead cells that leaked into the culture medium catalyzes the conversion of lactate to pyruvate and in the process generates NADH. In the presence of the diaphorase (reductase) (more...)
Other enzymes that do not use the NADH cycling assay chemistry also have been used as markers of dead cells. Examples include enzymes such as adenylate kinase (AK) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) that can produce ATP by providing a reaction cocktail containing the necessary ingredients to generate a cycling assay chemistry (8, 9); however, those enzymes may be less stable than LDH and lose enzymatic activity sooner after release from dead cells.
Another option is to measure protease activity as a marker that is released from dead cells with damaged membranes (10). Aminopeptidase activity can be measured using substrates containing a short sequence of amino acids (alanine-alanine-phenylalanine) conjugated via a peptide bond to either rhodamine 110 or aminoluciferin. Enzymatic removal of the amino acids can generate free rhodamine-110 for a fluorescent assay or free aminoluciferin which can be used by firefly luciferase to generate light (11).
Another optional assay approach is to load cells to contain a measurable marker. Loading target cells with an artificial measurable marker such as pro-fluorescent Calcein-AM (12) or radioactive 51Cr (13) has been used to measure cytotoxicity, mostly for assays that involve mixtures of more than one cell type (e.g. effector and target cells in antibody dependent cell mediated cytotoxicity assays). Target cells incubated with 51Cr will take up the radioactive marker which becomes bound as protein complexes in the cytoplasm of live cells. Similarly, calcein-AM is taken up by live cells where cytoplasmic esterase activity removes the AM group to generate fluorescent calcein which is retained in live cells. Upon cell death and loss of membrane integrity, the fluorescent calcein or the radioactive 51Cr is released from the cytoplasm into the culture medium where they can be quantified relative to the background spontaneous release from the viable cell population. The disadvantages of this approach include: the extra handling step to label the target cells prior to performing an assay, spontaneous release from the live cell population, and the safety and cost issues associated with using and disposing of radioactivity.
Still another cytotoxicity assay option is to genetically engineer cells to express luciferase as a marker (14). When cells die, the luminescence declines because the luciferase activity diminishes once cytoplasmic components are released into the culture medium. An advantage of this approach is the ability to selectively measure death of one type of cell in a mixed cell culture model such as antibody dependent cell mediated cytotoxicity assays. Disadvantages include the need to engineer cells to express luciferase and the assay measures a decrease in luminescent signal with increased cell killing which may make it difficult to detect small changes in the number of dead cells.
Commercial Availability
Calcein-AM; ThermoFisher Scientific Catalog No. C1430
CytoTox 96 Non-Radioactive Cytotoxicity Assay; Promega Corporation Catalog No. G1780
CytoTox-ONE Homogeneous Membrane Integrity Assay; Promega Corporation Catalog No. G7890
LDH-Glo Cytotoxicity Assay; Promega Corporation Catalog No. J2380.
CytoTox-Glo Cytotoxicity Assay; Promega Corporation Catalog No. G9290
Cytotoxicity Detection Kit (LDH); Millipore-Sigma (Roche) Catalog No. 11644793001
In Vitro Toxicology Assay Kit, Lactic Dehydrogenase based; Sigma Catalog No. TOX7-1KT
Pierce LDH Cytotoxicity Assay Kit; ThermoFisher Scientific Catalog No. 88953
aCella-TOX Glyceraldehyde-3 Phosphate Dehydrogenase; Cell Technology Catalog No. CLATOX100-3
Vybrant Cytotoxicity Assay Kit (Glucose-6-Phosphate Dehydrogenase Release Assay); ThermoFisher Scientific Catalog No. V23111
Example Protocol / Homogeneous Fluorescent LDH-Release Assay
The following example protocol is based on using the CytoTox-ONE Homogeneous Membrane Integrity Assay. For a detailed description of protocols including: reagent preparation, determination of linear range, detection sensitivity, and a detailed troubleshooting section, refer to Promega Technical Bulletin #306.
- 1.
Prepare opaque-walled assay plates containing cells in culture medium.
- 2.
Include control wells for: no cells (medium only), no treatment (cells with same amount of vehicle used to deliver test compounds), and maximum LDH release controls from detergent lysed cells (to determine value for 100% cytotoxicity).
- 3.
Add test compounds and vehicle controls to appropriate wells.
- 4.
Culture cells for desired test exposure period (usually 24-72 hours).
- 5.
Remove assay plates from 37°C incubator and equilibrate to 22°C (approximately 20–30 minutes) so all wells reach the same temperature.
- 6.
Add 2 μl of Lysis Solution (9% w/v Triton X-100) per 100 μl volume to the wells prepared for maximum LDH release control. (Note: If a larger pipetting volume is desired, use 10 μl of a 1:5 dilution of Lysis Solution.)
- 7.
Add a volume of CytoTox-ONE Reagent equal to the volume of culture medium present in each well.
- 8.
Mix for 30 seconds using a plate shaker.
- 9.
Incubate at 22°C for 10 minutes.
- 10.
Add Stop Solution (50 μl per 100 μl of CytoTox-ONE Reagent added) to each well. (Note: This step is optional, but recommended to stabilize the fluorescent signal and provide better consistency among replicate samples.)
- 11.
Mix the contents of the wells for 10 seconds using an orbital shaker and record resorufin fluorescence using an excitation wavelength of 560nm and an emission wavelength of 590nm.
shows example data from the LDH-release (CytoTox-ONE) assay. The effect of TNFα treatment on murine L929 cells was measured using the homogeneous (add-mix-measure) fluorescent LDH-release assay and a luminescent ATP assay as orthogonal methods using different endpoints to measure cell health. The half maximal response values correlate well for the LDH-release (dead cell) and the ATP content (viable cell number) assays.
Murine L929 cells were seeded at 2,000 cells per well in a 384-well opaque walled plate in serum-supplemented medium, cultured for 24 hours, then various amounts of TNFα were added and incubated overnight. Either CytoTox-ONE Reagent (to measure (more...)
Optional Protocol to Assist with Multiplexing
For assays that measure markers such as LDH released into the culture medium, an optional approach is to transfer an aliquot of culture supernatant into a separate suitable multi-well plate to perform the assay. Although this approach is a non-homogeneous protocol for the LDH-release assay, the advantage is that the population of cells remaining in the original assay plate can be used for multiplexing with another assay. The sample transfer approach can enable multiplexing in situations where two assay chemistries are not compatible or there is spectral overlap between fluorophores used for two different assays. Splitting the sample also has the result of reducing the volume of medium in the original well which may be required for multiplexing with assay reagents that recommend adding a 1:1 vol:vol ratio of reagent to culture medium. The extreme detection sensitivity of the luminescent version of the LDH-release assay provides an additional option of repeated sampling of 2-5 µl from the test wells at several different times to enable gathering kinetic data on the progress of cell death, while leaving the original sample of cells intact.
Controls for LDH-release Experiments
A negative control (background fluorescence) should be determined by adding the LDH detection reagent to replicate wells containing culture medium without cells and the same vehicle used to deliver test compound to experimental wells.
A 100% cell lysis positive control is recommended to determine the maximum amount of LDH present. Cells can be lysed to release LDH using an appropriate amount of a compound known to be toxic to the type of cells used in the experiment or a detergent. The CytoTox-ONE Homogeneous Membrane Integrity Assay described in the protocol above includes a Lysis Solution, which is a 9% (weight/volume) solution of Triton X-100 in water.
Troubleshooting
Animal sera used to supplement culture medium contains endogenous LDH activity and will contribute to background signal. Different lots and different types of sera contain different amounts of LDH activity. Using serum-free medium, reducing the percent of serum, or choosing a different lot or type can reduce background values.
The apparent half-life of LDH released from cells into the surrounding medium is approximately 9 hours; however, the half-life may depend on the mechanism of cell death and should be confirmed for each cell culture model used. If treated cells die (or if Lysis Solution is added) at the beginning of an experimental exposure period, the quantity of active LDH remaining in the culture medium at the end of the experiment may underestimate the quantity of LDH present in untreated cells. Performing 100% lysis of a set of control wells at the beginning and end of the incubation period is recommended. A further complication that should be considered is the amount of cell growth during the experiment. If the indicator cells are growing throughout the duration of the experiment, untreated control wells will have more cells and thus will have more LDH activity present at the end of the exposure period.
Pyruvate-supplemented media such as Ham’s F12, Iscove’s, and some formulations of DMEM may cause a reduction in the fluorescent signal due to product inhibition of the LDH enzymatic reaction catalyzing conversion of lactate to pyruvate.
The presence of strong reducing compounds may interfere with the resazurin-resorufin assay chemistry.
Measuring death of a specific population of cells can be challenging when there is more than one cell type present. Although LDH-release has been used for detecting target cell killing in mixtures of target and effector cells, additional controls must be used to account for spontaneous release of LDH from dying effector cells. This can be especially cumbersome to implement when several different ratios of effector to target cells are used for experiments.
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