Disadvantages of Metabolic Reduction Assays

Some of the major disadvantages of both the tetrazolium and resazurin reduction assays are related to the requirement to incubate a substrate with viable cells for a sufficient time to generate a measurable signal. This is a very important feature to consider when designing assays for HTS. In addition to the extra plate handling steps needed to return the cells to a 37°C incubator, the extended incubation of the detection reagents with viable cells increases the possibility of undesirable artifacts resulting from chemical interactions among the assay chemistries, the compounds tested, and the biochemistry of the cells.

One example is the known interference by reducing compounds that affect the chemical conversion of substrate to a colored indicator. This is especially true for the tetrazolium assays (Ulukaya, Colakogullari, and Wood 2004; Chakrabarti et al. 2000; Pagliacci et al. 1993; Collier and Pritsos 2003). The growing list of interfering compounds includes ascorbic acid and sulfhydryl reagents such as glutathione, coenzyme A, dithiothreitol, etc. Similar interferences by compounds that affect the oxidation and reduction chemistry of cells are likely to cause artifacts with the resazurin reduction assay. Assays that measure markers of metabolism also can be influenced by the pH of the culture medium and other factors that may stimulate or stress the metabolic rates of cells.

One of the major disadvantages of the tetrazolium and resazurin assays often overlooked is the toxic effects of the detection reagents on cells (Squatrito, Connor, and Buller 1995). Exposing cells to resazurin or tetrazolium reagents for long periods (or elevated concentrations for shorter periods) will result in cytotoxicity that has the potential to mask or interfere with the experimental outcome. The concentration of reagent and incubation time must be optimized to reduce cytotoxic effects to ensure avoidance of artifacts. Determining toxic effects can be accomplished by treating a population of cells with resazurin for various durations followed by determining viability using a different method such as measuring ATP. Figure 6.4 shows the toxic effects of Alamar blue on cells measured using ATP as a marker of viability.

Additional general limitations of all cell-based assays that utilize fluorescence detection methods include the possibility of fluorescence interference from small molecule compounds in chemical libraries and color quenching of signals from multiplex assays. Fluorescence interference can occur if the compound tested has properties that alter the normal detection of signal. Compounds may contribute to an artificially high fluorescent signal or may serve to quench normal signals from fluorescent indicators. Depending on the nature and size of the library tested, the effects of fluorescence interference may result in a substantial increase in the number of repeat assays to confirm hits or rule out interfering chemicals. The concentration of resazurin used for cell viability assays

70,000

60,000

oj 40,000

20,000 10,000 0

Resazurin

GF-AFC

Control

FIGURE 6.4 Comparison of effects of Alamar blue (resazurin) and GF-AFC reagents on viability of cells measured using a luminescent ATP assay. Resazurin or GF-AFC was incubated with 10,000 DU145 cells per well for 18 hr prior to measuring ATP as an indicator of cell viability. Alamar blue reagent is more toxic to cells.

produces solutions that are intensely colored and can result in quenching of secondary signals from other multiplexed fluorescent or luminescent assays. Despite several disadvantages that may lead to false hits and the need to repeat, the lower reagent cost per well has often been a deciding factor for choosing an assay method.

6.3.3.4 Aminopeptidase Markers

The recent identification of selective protease substrates to simultaneously measure markers of both viable and dead cells led to the development of optional methods for HTS that provide flexibility and added advantages (Niles et al. 2007a). The assay to measure viable cells is based on a cell-permeable protease substrate called glycyl-phenylalanyl-aminofluorocoumarin (GF-AFC). The procedure is a homogeneous add-incubate-measure method that is faster, more sensitive, and less toxic to cells than the tetrazolium and resazurin reduction assays. The substrate can be prepared in an aqueous buffer and is added directly to samples containing cells. The substrate permeates viable cells where constitutive protease activity in the cytoplasm rapidly removes the amino acids, yielding free AFC. The amount of AFC released is directly proportional to viable cell numbers and shows improved sensitivity compared to the resazurin assay (Figure 6.5). The AFC is detected via a microplate fluo-rometer equipped with a (380- to 400-nm excitation/505-nm emission) filter set.

The selective detection of viable cells by this method arises from the rapid loss of the protease activity that cleaves GF-AFC upon cell death, so only the viable population of cells contributes substantially to generating signals from free AFC. A 30-min incubation of the protease substrate with viable cells is generally sufficient to generate adequate signal for use in 384- or 1536-well format for HTS. The single liquid handling step contributes to improved assay performance and acceptable Z' factor values (Figure 6.6).

A major advantage of this method is the ability to multiplex with other assays. The GF-AFC substrate used to detect viable cells was designed for use in combination with another substrate that selectively detects protease activity from dead cells (Niles, Moravec, and Riss 2008). The method used to measure dead cells is based on the bis-Ala-Ala-Phe-rhodamine 110 (AAF-R110) protease substrate. This substrate is non-permeable; thus viable cells do not substantially contribute to signal. Dead cells with compromised membranes leak protease activity into the surrounding medium

10 100 1,000 10,000 100,000

Viable Cells/Well

10 100 1,000 10,000 100,000

Viable Cells/Well

FIGURE 6.5 Comparison of sensitivity of resazurin reduction and GF-AFC cleavage assays for detection of numbers of cells. Values are plotted as signal-to-noise (S:N) ratios. Comparison of data at S:N = 3 indicates approximately 30-fold better sensitivity of the GF-AFC assay after 30 min of incubation compared to resazurin incubated for 3.5 hr.

100,000

5 10,000 3

lu 1,000

R110 interference

No AFC interference

Color quenching

32 48

Well Number

FIGURE 6.6 Multiplexing cell viability and cytotoxicity assays to identify artifacts from various sources. HeLa cells (10,000 per well in 50 |L medium) were added to white, clear bottomed plates. Test compounds or controls diluted in 50 |L medium were added to appropriate wells. MultiTox-Fluor reagent (containing GF-AFC and AAF-R110) was prepared, added in 100 |L, and incubated at 370C for 30 min. Fluorescence was measured at 400 nm excitation/505 nm emission and 485 nm excitation/520 nm emission. Wells 1 to 4 were untreated viability controls. Wells 5 to 8 contained 30 |g/mL digitonin as a positive control to kill cells. Well 26 shows that R110 fluorescence interference at 485/520 scored as cytotoxic whereas the AFC fluorescence in well 26 was unaffected. The opposite scenario was demonstrated in well 79 where AFC fluorescence at 400/505 suggests an increase in viability. Wells 55, 57, and 75 containing cytotoxic compounds demonstrated an inverse relationship between the two biomarker values (long double-headed arrows). Well 88 demonstrated "flagging" of color quenching compounds not possible by a single parameter measure. Wells 54 and 63 received 5,000 and 20,000 cells, respectively, to demonstrate effects on magnitude of fluorescent values and how ratiometric measurements can be used to identify pipetting errors. (Source: Modified from Niles, A.L. et al. 2007b. Using protease biomarkers to measure viability and cytotoxicity. Promega Cell Notes 19, 16-20.)

where it can cleave amino acids from the AAF-R110 substrate and generate fluorescent rhodamine 110. The protease activity released from dead cells that cleaves AAF-R110 is relatively stable in culture medium, with a half-life of ~9 to 10 hr. The combination of the non-permeable nature of AAF-R110 and the rapid loss of the protease activity that cleaves GF-AFC upon cell death results in the ability to selectively detect both viable and dead populations of cells in the same sample.

The multiplex protocol to measure viable and dead cells in one sample utilizes a single reagent solution containing both the GF-AFC and AAF-R110 substrates. The combined substrates are added directly to samples of cells, incubated for 30 min, and fluorescence is recorded using two different filter sets to quantify the protease markers from the viable and dead populations. This homogeneous add-incubate-measure protocol to measure two endpoints can be used as an internal normalization control or provide additional useful information about the status of the population of cells treated (Figure 6.6). For example, if a treatment results in a decrease in viability measured with the GF-AFC substrate, you would expect to see an increase in dead cell marker activity in the same well. If fluorescence in the R110 channel increases (suggesting cell death) but the expected decrease in AFC fluorescence is not observed in that sample, the data may suggest that the tested compound did not reduce viable cell numbers but possibly contributed fluorescence in the R110 channel.

Measuring multiple parameters can be very useful for characterization of a cell culture model system during initial assay development and during subsequent optimization or miniaturization to a microwell plate format. The use of multiplexed internal controls also can be useful to identify false hits.

6.3.3.5 Luminogenic ATP Assay

Measurement of ATP is widely accepted as a valid method for estimating the number of viable cells from a variety of prokaryotic and eukaryotic sources. ATP levels are closely regulated by viable cells. When cells die, they lose the ability to synthesize ATP and endogenous ATPases rapidly deplete cytoplasmic stores to form ADP. ATP is a substrate for beetle luciferase. Although the chemical reaction is far more complex, a simplistic form to help explain the ATP assay for cell viability can be shown as:

ATP + luciferin + luciferase ^ light

Early protocols to measure ATP from cell cultures involved an acid or alkaline sample extraction step to precipitate proteins and inhibit ATPases (Lundin et al. 1986; Crouch et al. 1993). This was followed by removal of debris by centrifugation or filtration and a step to neutralize the pH of the sample prior to addition of luciferin and firefly luciferase to generate a flash of light proportional to the amount of ATP. Early reagents that created "flash" kinetics required luminometers with injection capabilities that hindered their use for HTS.

Years of effort have been dedicated to optimizing reagent systems to capitalize on the advantages of using luminescence as a signal in biological samples. Several technological breakthroughs and advances resulted in homogeneous procedures that require only a single reagent addition step to measure ATP in a variety of sample types. The elimination of the liquid handling steps provided flexibility to measure ATP in suspension or attached cell types.

Probably the most important advance enabling the development of the modern homogeneous ATP assay was the engineering of improved luciferase molecules. Directed evolution techniques were used to select luciferase mutants based on thermal stability (Hall et al. 1998). The result was identification of luciferases that could remain enzymatically active in the presence of ATPase inhibitors and harsh detergents used for cell lysis. These properties were critical for the development of robust ATP assays performed with a single reagent addition step and yielding extended signal stability.

The modern ATP assay has become the HTS method of choice to measure cell viability. It is the fastest and most sensitive assay and is less prone to artifacts than fluorescent methods. A growing list of publications (Rossi et al. 2007; Severson et al. 2007; Melnick et al. 2006) and examples posted to the PubChem Bioassays web site (http://pubchem.ncbi.nlm.nih.gov/assay/) make available data resulting from screening supported by the Roadmap Initiative of the National Institutes of Health.

The assay is performed by addition of a single reagent that contains a detergent to lyse cells immediately, ATPase inhibitors to stabilize the ATP present, and luciferin and luciferase contained in an optimized buffer formulation to allow the prolonged generation of photons of light. Within 10 min after the addition of reagent, the signal stabilizes and will glow for several hours, typically with a half-life of about 5 hr.

The ATP assay is faster because it uses a fundamentally different approach from tetrazolium and resazurin assay protocols that usually require 2- to 4-hr incubation when the plates must be returned to a 37°C incubator to allow viable cells to generate measurable product. The first step of the ATP assay includes cell lysis that eliminates co-incubation of the reagent and test compound in the presence of living cells. The homogeneous add-mix-measure protocol of the ATP assay also eliminates handling steps required to return assay plates to a cell culture incubator for development of a signal dependent on viable cells. In addition to making the assay more convenient, elimination of an incubation step reduces possible artifacts caused by interaction of the reagent chemistry with viable cells as occurs with tetrazolium and resazurin reduction assays.

The luminescent ATP assay is the most sensitive HTS method available for measuring the viability of cell populations in microwell plates. The limits of detection determined in samples of eukary-otic cells serially diluted from a known concentration may fall below 10 cells per well (Figure 6.7). This enables miniaturization to a 1536-well format.

Cells/Well

FIGURE 6.7 Sensitivity of detection of Jurkat cells using luminescent ATP assay. Serial twofold dilutions of cells were made in a 384-well solid white plate as 25 |L per well samples. An equal volume of ATP detection reagent was added and luminescence recorded after 10 min. Values represent mean + standard deviation of eight replicates. Direct relationship between luminescence and cell number (r2 = 0.99) is demonstrated. A Student's T-test indicates luminescence from a dilution containing four cells is significant over background. (Source: Modified from Hanna et al. 2001. CellTiter-Glo™ luminescent cell viability assay: a sensitive and rapid method for determining cell viability. Promega Cell Notes 2, 11-13.)

Cells/Well

FIGURE 6.7 Sensitivity of detection of Jurkat cells using luminescent ATP assay. Serial twofold dilutions of cells were made in a 384-well solid white plate as 25 |L per well samples. An equal volume of ATP detection reagent was added and luminescence recorded after 10 min. Values represent mean + standard deviation of eight replicates. Direct relationship between luminescence and cell number (r2 = 0.99) is demonstrated. A Student's T-test indicates luminescence from a dilution containing four cells is significant over background. (Source: Modified from Hanna et al. 2001. CellTiter-Glo™ luminescent cell viability assay: a sensitive and rapid method for determining cell viability. Promega Cell Notes 2, 11-13.)

Modification of the assay chemistry designed for measuring bacterial cells can achieve detection of lower concentrations of ATP (Junker and Clardy 2007; Fan et al. 2005). At this sensitivity, the assay performance becomes less limited by the ATP detection chemistry than the ability to repro-ducibly deliver and culture cells in microwell plates. The sensitivity is based on the high signal-to-background ratios that result from extremely low background luminescence. This is in contrast to fluorescent detection methods that require excitation of a fluorophore by illuminating a sample with an incident beam of light. The result is excitation of many other fluorescent molecules present in cells and serum supplemented culture media that can contribute to background fluorescence (Auld et al. 2008; Fan and Wood 2007).

6.3.3.6 Disadvantages of ATP Assay

One disadvantage of the ATP assay is the need for constant temperature to achieve consistent readings among different assay plates and within different locations within the same plate. The luminescent signal is dependent on the rate of the luciferase reaction that is related to temperature. For that reason, assay plates are usually allowed to equilibrate to a constant ambient temperature before addition of reagent. Complete equilibration is necessary because even slight temperature differences across the interiors, edges, and corners of a plate will result in altered luminescence readings.

With all assays that measure metabolic markers, cells with different metabolic capacities will produce different signal intensities. Factors that alter cell metabolism may influence the amount of ATP per cell and result in misleading data. The stimulation of the amount of ATP per cell has been used as an advantage in some applications such as measuring lymphocyte activation (Bulanova et al. 1995; White et al. 1989; Augustine, Pasi, and Hill 2007). Because ATP levels are tightly regulated, a decrease in ATP often results in cell death. Another disadvantage of the ATP assay (or any assay based on luciferase) is the possibility of interference by luciferase inhibitors present in small molecule chemical libraries. Molecules that interfere with luciferase measurements have been identified and annotated (Inglese et al. 2006; Kashem et al. 2007). Reagent formulations may affect the occurrence and extent of inhibition.

Many of these disadvantages can be overcome by implementing appropriate internal assay controls or by confirming results via an independent multiplexed measurement. One limitation of the ATP assay for multiplexing is that the protocol kills cells and produces a sample with high detergent and luciferin content. Sequential multiplexing of assays is still possible as long as the assay chemistries are compatible; however, other assays that use viable cells should be performed first in the sequence, before the addition of the ATP assay reagent. Examples of multiplexing with the ATP assay include genetic reporters, glutathione, cytochrome P450, resazurin reduction, GF-AFC aminopeptidase, etc.

6.4 APOPTOSIS ASSAYS

Programmed cell death was originally described based on morphological observations of developing tissues. Apoptosis is an active and well defined form of programmed cell death that plays an important role in regulating population dynamics of cells in multicellular organisms under normal and pathological conditions (Locksin and Zakeri 2001; Kerr, Wyllie, and Currie 1972; Wyllie, Kerr, and Currie 1980).

Apoptosis has served as an increasingly popular research topic over the past two decades because we recognize that elements in the apoptotic signaling pathways may be important therapeutic targets for treating cancer and other diseases. Interest in screening large libraries of small molecules has been growing in attempts to identify modulators of the apoptotic pathway and this led to the need for easy-to-use HTS assay methods.

A variety of methods are available to measure different processes occurring during apoptosis, including direct observation of morphological events, changes in mitochondrial membrane potential, movements of proteins to different subcellular compartments, DNA fragmentation, caspase-specific cleavage of target proteins, covalent binding and staining with fluorescent caspase inhibitors, and binding of fluorescently labeled Annexin V to phosphatidyl serine that becomes exposed on the outer leaflets of cell membranes.

Many elements of complex intrinsic and extrinsic signaling pathways leading to apoptosis have been identified. During apoptosis, a cascade of proteases termed caspases is involved with upstream signaling events and downstream executioner events. Caspases are cysteine-dependent, aspartate-spe-cific proteases that contain highly conserved cysteine residues in their active sites and cleave substrates leaving C terminal Asp residues. Caspase-3 is one of the main effector molecules of the apoptotic process. It cleaves several target proteins and serves as one of the executioner caspases that implement apoptosis. Despite reports of caspase-independent apoptosis (Broker et al. 2005), caspase-3 has become the most widely accepted and most frequently measured apoptosis marker for HTS.

Analogous to cell viability measurements, many of the same basic concepts apply to the development of cell-based assays for apoptosis. The length of incubation of cells with the test compound is among the most important issues to address and optimize. The length of incubation is important because the markers of apoptosis may be present for relatively brief transient periods and subsequently disappear as the population of cells undergoes secondary necrosis. The induction of measurable caspase activity can occur in only a few minutes or can take days, depending on the model cell line, type of inducer, and effective concentration inside the cells.

The kinetics of apoptosis induction depend on the specific cell type used and the culture environment, so it is advisable to optimize assay parameters for each cell line and also when changing culture conditions during miniaturization of assays.

To measure apoptosis, small peptide substrates selective for different caspase family members were developed based on technology used for other proteases (Gargiulo et al. 1981). Substrate design evolved from a colorimetric approach using paranitroanalide as the leaving group on the C terminal side of aspartic acid to fluorogenic substrates containing different coumarin derivatives or R110.

Although the peptide aminomethylcoumarin (AMC) substrate approach has been widely used for measuring the activities of many proteases, a disadvantage of this approach arises from the ultraviolet excitation and emission wavelengths required for detecting free AMC where there is a possibility for fluorescence interference. R110-labeled substrates are less encumbered by interference due to a spectral red shift, but many compounds in small molecule chemical libraries exhibit fluorescent properties that may interfere with assays based on fluorescence detection (Simenov et al. 2008).

Luminogenic substrates containing peptides linked to aminoluciferin overcome the problem of fluorescence interference while providing much better detection sensitivity (Monsees, Miska, and Geiger 1994; O'Brien et al. 2005). Figure 6.8 shows coupled enzymatic reactions in which caspase-3 cleaves the peptide substrate to liberate aminoluciferin that then becomes available as a substrate for the luciferase reaction to generate light. The luminogenic caspase-3 assay is the fastest and most sensitive method available for miniaturized HTS. The homogeneous add-incubate-read assay procedure involves adding a single reagent mixture directly to cells growing in multiwell plates. The samples are incubated for 30 to 60 min to achieve a steady state between the protease and luciferase enzymes, then luminescence is recorded to quantify the caspase-3 activity present. The reagent contains detergent, a stable form of luciferase, and ATP in a buffered formulation optimized to efficiently lyse cells and stabilize both caspase-3 and luciferase activities for hours. The half-life of the luminescent signal is typically >5 hr.

Direct comparison of the sensitivity of fluorogenic and luminogenic caspase-3 assays using R110 or aminoluciferin conjugated with the same DEVD peptide sequence has been reported to yield 20-fold greater sensitivity by the luminogenic method (O'Brien et al. 2005). As with any assay based on luciferase, the compounds may interfere with the luciferase activity. A decrease in luminescent signal may result from inhibition of caspase-3 or luciferase. A direct comparison of caspase-3 inhibition using fluorogenic and luminogenic assays to screen a library of 640 pharmacologically active compounds resulted in a lower false hit rate with the luminescent assay (O'Brien et al. 2005). Temperature is another factor that can affect the results of luminescent caspase-3 assays. To ensure consistent plate-to-plate results, samples are usually allowed to equilibrate to a constant ambient temperature before addition of reagent, similar to the procedure recommended for the ATP assay.

Z-DEVD-N

N COOH

Luciferase

Luciferase

N COOH

FIGURE 6.8 Dual enzymatic reactions of luminogenic protease assay to measure caspase-3 or -7 as an indicator of apoptosis. Caspase-3 ot -7 cleavage of the luminogenic protease substrate containing Z-DEVD-aminoluciferin releases aminoluciferin that can be used as a substrate for luciferase, resulting in production of light. (Source: Modified from Promega Corporation Technical Bulletin 323. Caspase-Glo® 3/7 Assay)

6.5 MULTIPLEXING CELL-BASED ASSAYS

False hits caused by assay artifacts represent a major concern for HTS. The risk of collecting flawed or misleading data can be partially mitigated by repeating an assay or using a confirmatory assay that detects a different parameter to measure the same event. Regardless of the type of cell-based assay used, knowing the number of viable cells at the end of the treatment period is practical information that can be utilized to normalize results. For example, if a single genetic reporter assay shows a decrease in signal, it is impossible to tell whether the decrease was a specific downregu-lation of the reporter or if the treatment led to cytotoxicity. Multiplexing both genetic reporter and viability assays in the same well provides additional insight that may suggest a mechanism of action (Figure 6.9). Multiplexing viability assays also can help reveal pipetting errors or differential growth patterns of cells across an assay plate.

To achieve multiplexing, the assay chemistries must be compatible and the signals must be distinguishable. Although it is desirable to measure both viability and apoptosis in the same sample, multiplexing a luminogenic ATP assay and a luminogenic caspase-3 assay is a combination that cannot be achieved because the reagent chemistries are not compatible. The reagent used to measure ATP contains an excess of luciferin—the compound detected in the luminogenic caspase-3 assay. The reagent used to measure caspase-3 activity contains an abundance of ATP—the marker measured in the luminescent viability assay. In cases such as this, multiplexing can be achieved by using appropriate assay reagents or by following a specified sequence of reagent additions. Figure 6.10 demonstrates that measuring viability and apoptosis from the same sample is possible using the GF-AFC protease substrate first in sequence to measure viable cell number followed by addition of the luminogenic caspase-3 reagent to quantify apoptosis (Niles, Moravec, and Riss 2008). The sequence of reagent addition is dictated by the parameter to be measured from a viable cell population. The GF-AFC protease substrate requires live cells, whereas the caspase-3 reagent lyses cells and thus must be added last.

CJ 3

ATP Indicator of Cell Viability

3,500

ATP Indicator of Cell Viability

3,500

CJ 3

Dual Reporter Assay Hours Iso

1 2 3 4 5 6 7 Hours After ISO/RO Induction

FIGURE 6.9 Sequential homogeneous multiplexing assay to measure Renilla luciferase reporter gene activity in live cells, followed by measuring ATP as an indicator of cell viability as a control in the same sample wells. Reporter gene activity in living cells increased over time, but cell numbers (ATP contents) remained constant. Multiplex data confirmed the increase in luminescence was due to specific upregulation of luciferase expression rather than an increase in viable cells. (Source: Modified from Riss, T.L. et al. 2005. Selecting cell-based assays for drug discovery screening. Promega Cell Notes 13, 16-21.)

1 2 3 4 5 6 7 Hours After ISO/RO Induction

FIGURE 6.9 Sequential homogeneous multiplexing assay to measure Renilla luciferase reporter gene activity in live cells, followed by measuring ATP as an indicator of cell viability as a control in the same sample wells. Reporter gene activity in living cells increased over time, but cell numbers (ATP contents) remained constant. Multiplex data confirmed the increase in luminescence was due to specific upregulation of luciferase expression rather than an increase in viable cells. (Source: Modified from Riss, T.L. et al. 2005. Selecting cell-based assays for drug discovery screening. Promega Cell Notes 13, 16-21.)

Another approach to overcome assay chemistry incompatibility is splitting a sample into two different containers. If the cell population releases a marker into the surrounding environment, a small aliquot of culture medium can be sampled and moved to a different assay plate to segregate assay chemistries. Figure 6.11 shows the measurement of lactate dehydrogenase released from a

i 10,000

• GF-AFC Cell viability assay O Caspase 3/7 apoptosis assay

• GF-AFC Cell viability assay O Caspase 3/7 apoptosis assay

Log10 [paclitaxel] M

r 6,000

4,000 g

Log10 [paclitaxel] M

FIGURE 6.10 Multiplexing of fluorescent cell viability assay measuring cleavage of GF-AFC with the lumi-nogenic apoptosis assay measuring caspase-3 cleavage of Z-DEVD-aminoluciferin. The GF-AFC reagent was added to wells containing 10,000 cells per well, incubated 30 min at 370C, and fluorescence measured as an indicator of cell viability. Luminogenic caspase-3/7 reagent was added, incubated 30 min at room temperature and luminescence measured as an indicator of apoptosis. The inverse relationship confirms loss of viability due to apoptosis. (Source: Modified from Promega Corporation Technical Bulletin 371. CellTiter-Fluor™ Cell Viability Assay.)

Cell Titer Fluor

FIGURE 6.11 Multiplex measurement of cytotoxicity and apoptosis markers from the same sample of cells accomplished by separating incompatible assay chemistries into different containers. HepG2 cells (10,000 per well) were plated in a solid white and clear bottom 96-well plate and cultured overnight. Various concentrations of tamoxifen were added to the wells and incubated for 4 hr at 370C. To assay LDH activity, 50 |L per well of culture supernatant was transferred to a 96-well plate to which 50 |L per well of LDH assay reagent was added. LDH samples were incubated at ambient temperature for 30 min prior to stopping the reaction to measure fluorescence at 560/590 nm. For caspase-3/7 determination, 50 |L per well of bis-DEVD-R110 reagent was added to the original culture plate containing cells, and incubated at ambient temperature for 45 min prior to determining fluorescence at 485/527 nm. (Source: Modified from Cell Viability Protocols and Application Guide. Madison, WI: Promega Corporation.)

FIGURE 6.11 Multiplex measurement of cytotoxicity and apoptosis markers from the same sample of cells accomplished by separating incompatible assay chemistries into different containers. HepG2 cells (10,000 per well) were plated in a solid white and clear bottom 96-well plate and cultured overnight. Various concentrations of tamoxifen were added to the wells and incubated for 4 hr at 370C. To assay LDH activity, 50 |L per well of culture supernatant was transferred to a 96-well plate to which 50 |L per well of LDH assay reagent was added. LDH samples were incubated at ambient temperature for 30 min prior to stopping the reaction to measure fluorescence at 560/590 nm. For caspase-3/7 determination, 50 |L per well of bis-DEVD-R110 reagent was added to the original culture plate containing cells, and incubated at ambient temperature for 45 min prior to determining fluorescence at 485/527 nm. (Source: Modified from Cell Viability Protocols and Application Guide. Madison, WI: Promega Corporation.)

population of tamoxifen-treated necrotic cells (Korzeniewski and Callewaert 1983). An aliquot of culture medium was removed to a separate assay plate without disturbing the remaining cells in the original plate that was then treated with a cell lysis reagent containing a fluorogenic peptide substrate to measure caspase-3 activity as a marker of apoptosis. Although this multiplexing approach is not considered homogeneous because of a liquid transfer step, it still presents the advantage of providing two different measurements from the same population of cells.

6.6 SUMMARY AND CONCLUSIONS

A variety of assay technologies have been developed to enable miniaturized screening of viability and apoptosis. Each approach has advantages and disadvantages that must be considered before choosing the most appropriate assay. The ultimate goal is to obtain the highest quality data set in the shortest time by the most cost-effective method possible.

It is important the assay technology does not lead to artifacts caused by interactions of the reagent chemistry with physiological events in the cells. Although the tetrazolium and resazurin reduction assays historically gained popularity because they are homogeneous and do not require the removal of culture medium from the sample, the limited detection sensitivity and the need to incubate viable cells with reagent for 1 to 4 hr to generate an adequate signal are among the reasons the ATP viability assay has become the most popular for HTS. Similar advantages of speed and improved sensitivity to enable miniaturization led to the current popularity of the luminogenic caspase assay approach for detecting apoptosis in HTS.

Understanding the biology and kinetics of the cell death process and realizing that cell culture is an artificial model system can help guide characterization experiments to develop the best assay possible. Probably the most common problem facing design of viability and apoptosis assays for HTS is deciding on the duration of incubation with the test compounds before making a measurement.

Often the results of screening campaigns from pharmaceutical or biotechnology companies are not published to protect their intellectual property positions; however, a growing collection of screening data from NIH-funded programs is now available to the public (including descriptions of methods used) through the PubChem bioassay web site. These postings provide insight into the types of assays used in the academic screening community. The assays posted have undergone sufficient development to verify their readiness for HTS. Additional resources are available to aid in assay development, including an Assay Guidance Manual resulting from a collaboration between Eli Lilly & Co. and NIH's Chemical Genomics Center. This resource is currently available on the Internet and continues to be expanded to cover more topics.

The weaknesses of some assay approaches can be compensated for during counter screening or by simultaneously multiplexing with a secondary assay. Options are available for multiplexing viability and apoptosis assays with each other or as internal normalization controls for other cell-based assays. The ability to multiplex assay measurements in entire populations of cells can be an efficient substitute for a high content imaging approach, especially when screening a large collection of compounds or when automated imaging instrumentation is not available.

REFERENCES

Ahmed, S.A., Gogal, R.M., and Walsh, J.E. 1994. A new rapid and simple nonradioactive assay to monitor and determine the proliferation of lymphocytes: an alternative to [3H]thymidine incorporation assays. J. Immunol. Meth. 170. 211-224. Augustine, N.H., Pasi, B.M., and Hill, H.R. 2007. Comparison of ATP production in whole blood and lymphocyte proliferation in response to phytohemagglutinin. J. Clin. Lab. Anal. 21, 265-270. Auld, D.S. et al. 2008. Characterization of chemical libraries for luciferase inhibitory activity. J. Med. Chem. 51, 2372-2386.

Barltrop, J. and Owen, T. 1991. 5-(3-carboxymethoxyphenyl)-2-(4,5-dimethylthiazoly)-3-(4-sulfophenyl)tetra-zolium, inner salt (MTS) and related analogs of 3-(4,5-dimethylthiazolyl)-2,5-diphenyltetrazolium bromide (MTT) reducing to purple water-soluble formazans as cell viability indicators. Bioorg. Med. Chem. Lett. 1, 611-614.

Borenfreund, E. and Puerner, J.A. 1985. Toxicity determined in vitro by morphological alterations and neutral red absorption. Toxicol. Lett. 24, 119-124. Bröker, L.E. et al. 2005. Cell death independent of caspases: a review. Clin. Cancer Res. 11, 3155-3162. Bulanova, E.G. et al. 1995. Bioluminescent assay for human lymphocyte blast transformation. Immunol. Lett. 46, 153-155.

Cavanaugh, P.F. et al. 1990. A semi-automated neutral red based chemosensitivity assay for drug screening.

Invest. New Drugs 8, 347-354. Chakrabarti, R. et al. 2000. Vitamin A as an enzyme that catalyzes the reduction of MTT to formazan by vitamin C. J. Cellular Biochem. 80, 133-138. Chatterjee, R. 2007. Cases of mistaken identity. Science 315, 928-930.

Collier, A. and Pritsos, C. 2003. The mitochondrial uncoupler dicumerol disrupts the MTT assay. Biochem. Pharm. 66, 281-287.

Cory, A. et al. 1991. Use of an aqueous soluble tetrazolium/formazan assay for cell growth assays in culture.

Cancer Commun 3, 207-212. Crouch, S.P. et al. 1993. The use of ATP bioluminescence as a measure of cell proliferation and cytotoxicity.

J. Immunol. Meth. 160, 81-88. Denizot, F. and Lang, R. 1986. Rapid colorimetric assay for cell growth and survival: modifications to the tetrazolium dye procedure giving improved sensitivity and reliability. J. Immunol. Meth. 89, 271-277.

Digan, M.E. et al. 2005. Evaluation of division-arrested cells for cell-based high-throughput screening profiling. J. Biomol. Screen. 10, 615-623. Ekwall, B. et al. 1998. MEIC evaluation of acute systemic toxicity VI: prediction of human toxicity by rodent LD50 values and results from 61 in vitro methods. ATLA 26, 617-658.

Essodaigui, M., Broxterman. H.J., and Garnier-Suillerot, A. 1998. Kinetic analysis of calcein and calcein-acetoxymethylester efflux mediated by the multidrug resistance protein and P-glycoprotein. Biochemistry 37, 2243-2250.

Fan, F. et al. 2005. BacTiter-Glo™ assay for antimicrobial drug discovery and general microbiology. Promega Notes 89, 25-27.

Fan, F. and Wood, K. 2007. Bioluminescent assays for high-throughput screening. Assay Drug Dev. Technol. 5:127-136.

Gargiulo, R.J. et al. 1981. Analytical fluorogenic substrates for proteolytic enzymes. United States Patent 4275153.

Garner, D.L. et al. 1994. Dual DNA staining assessment of bovine sperm viability using SYBR-14 and pro-pidium iodide. J. Androl. 15, 620-629.

Hall, M.P. et al. 1998. Stabilization of firefly luciferase using directed evolution. In Bioluminescence and Chemiluminescence: Perspectives for the 21st Century, Roda, A., et al., Eds. Chichester: John Wiley & Sons, 392-395.

Hanna et al. 2001 CellTiter-G10™ luminescent cell viability assay: a sensitive and rapid method for determining cell viability. Promega Cell Notes 2, 11-13.

Hansen, M.B., Nielsen, S.E. and Berg, K. 1989. Re-examination and further development of a precise and rapid dye method for measuring cell growth/cell kill. J. Immunol. Meth. 119, 203-210.

Inglese, J. 2008. Characterization of chemical libraries for luciferase inhibitory activity. J. Med. Chem. 51, 2372-2386.

Inglese, J. et al. 2006. Quantitative high-throughput screening: a titration-based approach that efficiently identifies biological activities in large chemical libraries. Proc. Natl. Acad. Sci. USA 103, 11473-11478.

Ishiyama, M. et al. 1993. A new sulfonated tetrazolium salt that produces a highly water-soluble formazan dye. Chem. Pharm. Bull. 41, 1118-1122.

Ivnitski-Steele, I. et al. 2008. High-throughput flow cytometry to detect selective inhibitors of ABCB1, ABCC1, and ABCG2 transporters. Assay Drug. Dev. Technol. 6, 263-276.

Junker, L.M. and Clardy, J. 2007. High-throughput screens for small-molecule inhibitors of Pseudomonas aeruginosa biofilm development. Antimicrob. Agents Chemother. 51, 3582-3590.

Karaszi, E. et al. 2001. Calcein assay for multidrug resistance reliably predicts therapy response and survival rate in acute myeloid leukaemia. Br. J. Haematol. 112, 308-314.

Kashem. M. et al. 2007. Three mechanistically distinct kinase assays compared: measurement of intrinsic ATPase activity identified the most comprehensive set of ITK inhibitors. J. Biomol. Screen. 12, 70-83.

Kerr, J.F., Wyllie, A.H., and Currie, A.R. 1972. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br. J. Cancer 26, 239-257.

Korzeniewski, C. and Callewaert, D.M. 1983. An enzyme-release assay for natural cytotoxicity. J. Immunol. Meth. 64, 313-320.

Lockshin, R.A. and Zakeri, Z. 2001. Programmed cell death and apoptosis: origins of the theory. Nat. Rev. Mol. Cell Biol. 2, 545-550.

Lundin, A. et al. 1986. Estimation of biomass in growing cell lines by ATP assay. Meth. Enzymol. 133, 27-42.

Melnick, J.S. et al. 2006. An efficient rapid system for profiling the cellular activities of molecular libraries. Proc. Natl. Acad. Sci. USA 103, 3153-3158.

Monsees, T., Miska, W. and Geiger, R. 1994. Synthesis and characterization of a bioluminogenic substrate for a-chymotrypsin. Anal. Biochem. 221, 329-334.

Mosmann, T. 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol. Meth. 65, 55-63.

Niles, A., Moravec, R., and Hesselberth, E. 2007. A homogeneous assay to measure live and dead cells in the same sample by detecting different protease markers. Anal. Biochem. 366, 197-206.

Niles, A.L. et al. 2007b. Using protease biomarkers to measure validity and cytotoxicity. Promega Cell Notes 19, 16-20.

Niles, A., Moravec, R., and Riss, T.L. 2008. Multiplex caspase activity and cytotoxicity assays. Methods Mol. Biol. 414, 151-162.

O'Brien, M. et al. 2005. Homogeneous, bioluminescent protease assays: caspase-3 as a model. J. Biomol. Screen. 10, 137-148.

O'Brien, P. and Haskins, J.R. 2006. In vitro cytotoxicity assessment. In Methods in Molecular Biology 356: High Content Screening Taylor, D.L. et al. Totowa, NJ: Humana, 415-425.

Pagliacci, M. et al. 1993. Genistein inhibits tumour cell growth in vitro but enhances mitochondrial reduction of tetrazolium salts: a further pitfall in the use of the MTT assay for evaluating cell growth and survival. Eur. J. Cancer 29, 1573-1577.

Paull, K.D. et al. 1988. The synthesis of XTT: a new tetrazolium reagent that is bioreducible to a water-soluble formazan. J. Heterocyclic Chem. 25, 911-914.

Riss, T.L. et al., 2005. Selecting cell-based assays for drug discovery screening. Promega Cell Notes 13, 16-21.

Riss, T.L. and Moravec, R.A. 2004. Use of multiple assay endpoints to investigate the effects on incubation time, dose of toxin, and plating density in cell-based cytotoxicity assays. Assay Drug Dev. Technol. 2, 51-62.

Rossi, C. et al. 2007. Identifying druglike inhibitors of myelin-reactive T cells by phenotypic high-throughput screening of a small-molecule library. J. Biomol. Screen. 12, 481-489.

Rubinstein, L.V. et al. 1990. Comparison of in vitro anticancer drug screening data generated with a tetrazolium assay versus a protein assay against a diverse panel of human tumor cell lines. J. Natl. Cancer Inst. 82, 1113-1117.

Severson, W.E. et al. 2007. Development and validation of a high-throughput screen for inhibitors of SARS CoV and its application in screening of a 100,0000-compound library. J. Biomol. Screen. 12, 33-40.

Shoemaker, R.H. 2006. The NCI60 human tumour cell line anticancer drug screen. Nat. Rev. Cancer 6, 813-823.

Shum, D. et al. 2008. A high density assay format for the detection of novel cytotoxic agents in large chemical libraries. J. Enz. Inhib. Med. Chem. 23, 931-945.

Simenov, A. et al. 2008. Fluorescence spectroscopic profiling of compound libraries. J. Med. Chem. 51, 2363-2371.

Skehan P. 1990. New colorimetric cytotoxicity assay for anticancer-drug screening. J. Natl. Cancer Inst. 82, 1107-1112.

Skehan, P. et al. 1989. Evaluation of colorimetric protein and biomass stains for assaying drug effects upon human tumor cell lines. Proc. Amer. Assoc. Cancer Res. 30, 612-616.

Squatrito, R., Connor, J., and Buller, R. 1995. Comparison of a novel redox dye cell growth assay to the ATP bioluminescence assay. Gynecol. Oncol. 58, 101-105.

Tada, H. et al. 1986. An improved colorimetric assay for interleukin 2. J. Immunol. Meth. 93, 157-165.

Tominaga, H. et al. 1999. A water-soluble tetrazolium salt useful for colorimetric cell viability assay. Anal. Commun. 36, 47-50.

Ulukaya, E., Colakogullari, M., and Wood, E.J. 2004. Interference by anti-cancer chemotherapeutic agents in the MTT-tumor chemosensitivity assay. Chemotherapy 50, 43-50.

Vukicevic, S. et al. 1992. Identification of multiple active growth factors in basement membrane Matrigel suggests caution in interpretation of cellular activity related to extracellular activity related to extracellular matrix components. Exp. Cell Res. 202, 1-8.

Wesierska-Gadek, J. et al. 2005. A new multiplex assay allowing simultaneous detection of the inhibition of cell proliferation and induction of cell death. J. Cell. Biochem. 96, 1-7.

White, A.G. et al. 1989. Lymphocyte activation: changes in intracellular adenosine triphosphate and deoxyribonucleic acid synthesis. Immunol Lett. 22, 47-50.

Wigglesworth, K.J. et al. 2008. Use of cryopreserved cells for enabling greater flexibility in compound profiling. J. Biomol. Screen. 13, 354-362.

Wyllie, A., Kerr, J., and Currie, A. 1980. Cell death: the significance of apoptosis. Int. Rev. Cytol. 68, 251-306.

Xia, M. et al. 2008. Compound cytotoxicity profiling using quantitative high-throughput screening. Envir. Health Perspect. 116, 284-291.

Zaman, G.J.R. et al. 2007. Cryopreserved cells facilitate cell-based drug discovery. Drug Disc. Today 12, 521-526.

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  • kari
    What does resazurin reduction assay measure?
    2 months ago

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