update on in vitro cytotoxicity assays for drug development

16
Review 10.1517/17460440802163960 © 2008 Informa UK Ltd ISSN 1746-0441 655 Update on in vitro cytotoxicity assays for drug development Andrew L Niles , Richard A Moravec & Terry L Riss Promega Corporation, Research and Development, 2800 Woods Hollow Road, Madison, Wisconsin, 53711, USA Background: In vitro cytotoxicity testing provides a crucial means of ranking compounds for consideration in drug discovery. The choice of using a particular viability or cytotoxicity assay technology may be influenced by specific research goals. Objective: Although the high-throughput screening (HTS) utility is typically dependent upon sensitivity and scalability, it is also impacted by signal robustness and resiliency to assay interferences. Further consideration should be given to data quality, ease-of-use, reagent stability, and matters of cost-effectiveness. Methods: Here we focus on three main classes of assays that are at present the most popular, useful, and practical for HTS drug discovery efforts. These methods measure: i) viability by metabolism reductase activities; ii) viability by bioluminescent ATP assays; or iii) cytotoxicity by enzymes ‘released’ into culture medium. Multi-parametric technologies are also briefly discussed. Results/conclusion: Each of these methods has its relative merits and detractions; however multi-parametric methods using both viability and cytotoxicity markers may mitigate the inherent shortcomings of single parameter measures. Keywords: apoptosis, cytostasis, cytotoxicity, fluorescence, high-throughput, luminescence, multi-parametric, multiplex, necrosis, resazurin, tetrazolium, viability Expert Opin. Drug Discov. (2008) 3(6):655-669 1. Introduction The term ‘cytotoxicity’ often has a broad and ill-defined meaning in the drug discovery and development industry. For in vitro cell culture systems, a compound or treatment is considered to be cytotoxic if it interferes with cellular attachment, significantly alters morphology, adversely affects cell growth rate, or causes cell death [1]. A variety of assays have been developed and used for the measurement of viability or cytotoxicity in vitro, including classical dye inclusion or exclusion and colony formation assays [2-4]. Unfortunately many of these methods are laborious, time-consuming, insensitive, and thus poorly suited for high-throughout screening (HTS). To this end, more efficient modern screening methods have been devised that examine and measure cessation of a broad variety of parameters associated with biochemical events necessary for sustaining viability and/or evidence of changes in membrane integrity leading to cellular disintegration [5,6]. Depending on the ultimate goals of the screen, one must choose to use either a viability or a cytotoxicity assay, or some combination of both in conjunction with each other. This decision rests on the fact that there is a large practical distinction between the data derived from viability and cytotoxicity measures. The scientific premise behind metabolism and ATP-based viability assays is that activity is proportional to viable cell number. As such, it is strongly inferred that a reduction in activity after treatment (when compared with control) is the result of cytotoxicity. Conversely, cytotoxicity assays measure parameters proportional to the degree of cell death in an assay well. During most cytotoxicity events, viability and cytotoxicity measures are inversely proportional. That is, viability measures 1. Introduction 2. Main classes of viability or cytotoxicity assays 3. Conclusions 4. Expert opinion Expert Opin. Drug Discov. Downloaded from informahealthcare.com by Universita Studi di Torino on 05/06/13 For personal use only.

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Page 1: Update on               in vitro               cytotoxicity assays for drug development

Review

10.1517/17460440802163960 © 2008 Informa UK Ltd ISSN 1746-0441 655

Update on in vitro cytotoxicity assays for drug development Andrew L Niles † , Richard A Moravec & Terry L Riss Promega Corporation, Research and Development, 2800 Woods Hollow Road, Madison, Wisconsin, 53711, USA

Background : In vitro cytotoxicity testing provides a crucial means of ranking compounds for consideration in drug discovery. The choice of using a particular viability or cytotoxicity assay technology may be influenced by specific research goals. Objective : Although the high-throughput screening (HTS) utility is typically dependent upon sensitivity and scalability, it is also impacted by signal robustness and resiliency to assay interferences. Further consideration should be given to data quality, ease-of-use, reagent stability, and matters of cost-effectiveness. Methods : Here we focus on three main classes of assays that are at present the most popular, useful, and practical for HTS drug discovery efforts. These methods measure: i) viability by metabolism reductase activities; ii) viability by bioluminescent ATP assays; or iii) cytotoxicity by enzymes ‘released’ into culture medium. Multi-parametric technologies are also briefly discussed. Results/conclusion : Each of these methods has its relative merits and detractions; however multi-parametric methods using both viability and cytotoxicity markers may mitigate the inherent shortcomings of single parameter measures.

Keywords: apoptosis , cytostasis , cytotoxicity , fluorescence , high-throughput , luminescence , multi-parametric , multiplex , necrosis , resazurin , tetrazolium , viability

Expert Opin. Drug Discov. (2008) 3(6):655-669

1. Introduction

The term ‘cytotoxicity’ often has a broad and ill-defined meaning in the drug discovery and development industry. For in vitro cell culture systems, a compound or treatment is considered to be cytotoxic if it interferes with cellular attachment, significantly alters morphology, adversely affects cell growth rate, or causes cell death [1] . A variety of assays have been developed and used for the measurement of viability or cytotoxicity in vitro , including classical dye inclusion or exclusion and colony formation assays [2-4] . Unfortunately many of these methods are laborious, time-consuming, insensitive, and thus poorly suited for high-throughout screening (HTS). To this end, more efficient modern screening methods have been devised that examine and measure cessation of a broad variety of parameters associated with biochemical events necessary for sustaining viability and/or evidence of changes in membrane integrity leading to cellular disintegration [5,6] .

Depending on the ultimate goals of the screen, one must choose to use either a viability or a cytotoxicity assay, or some combination of both in conjunction with each other. This decision rests on the fact that there is a large practical distinction between the data derived from viability and cytotoxicity measures. The scientific premise behind metabolism and ATP-based viability assays is that activity is proportional to viable cell number. As such, it is strongly inferred that a reduction in activity after treatment (when compared with control) is the result of cytotoxicity. Conversely, cytotoxicity assays measure parameters proportional to the degree of cell death in an assay well. During most cytotoxicity events, viability and cytotoxicity measures are inversely proportional. That is, viability measures

1. Introduction

2. Main classes of viability

or cytotoxicity assays

3. Conclusions

4. Expert opinion

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Page 2: Update on               in vitro               cytotoxicity assays for drug development

Update on in vitro cytotoxicity assays for drug development

656 Expert Opin. Drug Discov. (2008) 3(6)

are high when cytotoxicity measures are low, and vice versa. However, the fundamental difference between the approaches becomes pronounced depending on the length of compound exposure [5] . For instance, short-term toxin exposures (4 h or less) may adversely affect metabolism markers or ATP content before measurable membrane integrity changes [7-9] . Long-term exposures (24 h or greater), particularly after early primary necrosis, may lead to underestimation of cytotoxicity owing to degradation of marker enzyme activity after release into the extracellular environment [5] .

In addition to considerations surrounding the kinetics of cell death, another caveat of preferentially using a viability

or cytotoxicity measure in lieu of both parameters comes in instances of cytostasis, or cell-cycle arrest. Most historically useful and therapeutically attractive anticancer compounds exhibit antiproliferative effects for sustained time periods (up to 48 h or more) before demonstrating cytotoxicity [10] . Therefore, prolonged cell-cycle arrest may lead to false negative results in a cytotoxicity assay or may lead to a false positive test result with a viability assay in cases where cells may ultimately relieve the block and escape stasis ( Figure 1 ).

Concentration dependence is another important aspect of cytotoxicity screening. Many cell-based screens of the

Viability EC50 = N.D.

Cytotoxicity EC50 = N.D.

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Log10 [paclitaxel] M

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Viability EC50 = 26.1 nM

Cytotoxicity EC50 = 25.4 nM

Log10 [nocodazole] M

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Viability EC50 = 423 nM

Cytotoxicity EC50 = N.D.

Log10 [doxorubicin] M

EC50 = 15 nM (partial fit)

Viability EC50 = 27.6 nM

Cytotoxicity EC50 = N.D.

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A. B.

C. D.

Figure 1 . Viability and cytotoxicity profi les can vary depending on compound potency, dosage, and time of exposure. Cells were exposed to compounds and then subjected to same-well viability/cytotoxicity chemistries which measure separate and distinct proteolytic biomarkers (MultiTox-Fluor or MultiTox-Glo, Promega Corporation). A. H929 cells demonstrate no deleterious effect after 24 h of paclitaxel exposure. B. K562 cells demonstrate concentration-dependent, cell-cycle arrest without cytotoxicity after 24 h of doxorubicin treatment. C. K562 cells show a reduction in viability with a commensurate increase in cytotoxicity after treatment with nocodazole. D. H929 cells display a concentration-dependent reduction in viability and a cytotoxicity profi le consistent with late-stage cytotoxicity after 24 h of colchicine treatment. The decline in the cytotoxicity biomarker activity at the highest concentration is due to concentration-dependent kinetics of cell death and degradation of the cytotoxicity biomarker as a function of time in the extracellular environment.

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1990s used single or replicate assay wells at a single compound concentration (e.g., 10 µM, final). This approach allows for testing of a greater number of chemical compounds but is problematic from a statistical viewpoint owing to biological variation in response and physiochemical concerns such as compound solubility [11] . The quantitative high-throughput screening (qHTS) approach championed by Inglese et al. involves examining each compound in a screen in broad serial dose dilutions [12] . This approach is initially more technically involved but is gaining scientific momentum because it produces high quality response curves that allow for greater characterization of cytotoxic effects while mitigating false positive or negative test results [13] .

The last main consideration in cytotoxicity testing is the manner in which cells die. The mechanism of death can be quite important if cytotoxicity testing is initiated for ancillary safety concerns (e.g., off-target cytotoxicity from pharmaceuticals, cosmetics, and nutritional supplements), or specifically, as for identifying new chemical entities for cancer therapy. Simply stated, compounds that cause primary necrosis in cell culture may carry unacceptable cytotoxic liabilities, whereas compounds that cause apoptosis can be preferable. Again, compound concentration and cell death kinetics may vary but generally primary necrosis or catastrophic cell lysis occurs very quickly after compound addition (2 h or less), whereas apoptosis proceeds in a more orderly manner (4 – 48 h) ( Figure 2 ) [14-16] .

Time zero 30 min to 4 h 4 to 48 h > 48 h

2° NecrosisViable cell

Apoptosis

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release0 +++++ +

Caspase 0 ++++++ 0

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Figure 2 . Chronological comparison of measurable viability and cytotoxicity biomarkers during typical apoptotic or primary necrotic events. Apoptosis causes a time-dependent (typically 4 – 48 h) reduction in viability marker activity with a commensurate increase in cytotoxicity marker activity. Primary necrosis presents a similar profi le, albeit in a much shorter time frame (typically 30 min to 4 h). Therefore, the kinetics of the cytotoxic response may infl uence which assay chemistry is most appropriate for a study.

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658 Expert Opin. Drug Discov. (2008) 3(6)

2. Main classes of viability or cytotoxicity assays

2.1 Viability assays based on metabolism reductase activities Many assay methods have been described over the past 25 years to estimate cell viability based on various cellular metabolism parameters, including those based on tetrazolium and resazurin reduction. Several tetrazoliums have evolved, with each one developed to improve on one or more limitations of the previous. For example, the original reference incorpo-rating the tetrazolium MTT (3-(4,5-dimethylthiazolyl)-2,5-diphenyltetrazolium bromide) is regarded as the first example of a tetrazolium salt that was used in the development of a multi-well viability assay for mammalian cells [17] . Originally described as a two-addition assay, it revolutionized cell-based drug screening by offering a HTS colorimetric assay that simplified sample processing and did not require radioisotope but was sensitive enough for miniaturization into 96-well plate formats. The procedure involves the addition of a small volume of MTT contained in phosphate-buffered saline to the cell culture. During 1 – 4 h of incubation at 37 ° C, a pool of cellular mitochondrial and cytosolic enzymes reduces the dye to its reduced purple formazan form [18] . Cellular reduction of MTT yields an aqueous insoluble formazan, with visible crystals contained both in the surrounding medium as well as within the cells. For this reason, a second addition containing an acid/isopropanol solvent is required to solubilize and disperse the formazan before recording absorbance values. As the assay procedure was further scrutinized, it became apparent that for many cell lines and protocols, the amount of formazan produced and the presence of culture medium serum warranted removal of the culture medium and addition of dimethyl sulfoxide (DMSO) to generate maximum absorbance [19] . DMSO and organic solvent based solubilization solutions exposed laboratory personnel and equipment to unnecessary chemical fumes and risk, and removal of the culture medium added a third step in sample processing [20] . In addition, by adding an organic solvent mixture to solubilize the formazan, the assay becomes a lytic end point assay, offering no opportunity for additional readings if further color development is needed, particularly important during assay optimization and characterization. Despite the early problematic issues surrounding solubilization, many different commercial reagents were developed, thereby improving this technology. The MTT assay is extremely well charac-terized and referenced to this day in the literature, and is often a gold standard that new viability/cytotoxicity assay methods are compared with.

Problems surrounding MTT’s insoluble formazan product prompted investigation and the creation of alternative tetrazoliums with different properties. XTT (2,3-bis(2-methoxy-4-nitro-5-sulphophenyl)-5-carboxanilide-2H-tetrazolium) was the first tetrazolium developed that yielded an aqueous

soluble formazan product upon cellular reduction [21] . The two sulfonic acid groups on XTT are critical for solubility as a formazan product and gave this viability assay a new edge by eliminating the need for a second addition of solubilization solvent. However, XTT has struggled to gain widespread acceptance owing to its inherent limited solubility in aqueous buffers. Furthermore, stock solutions proved to be difficult to prepare and had limited stability as a reagent [22] .

MTS (5-(3-carboxymethoxyphenyl)-2-(4,5-dimethylthiazoly)-3-(4-sulfophenyl)tetrazolium, inner salt) [23,24] and WST-1 ((4-[3-4-iodophenyl]-2-(4-nitrophenyl)-2H-5-tetrazolio)-1,3-benzene disulfonate) [25] were developed as third gene-ration tetrazoliums that exhibit good solubility, are easy to prepare, and are very stable as stock solutions. The chemical properties of XTT, MTS, and WST-1 tetrazoliums warrant that they be formulated with an electron-coupling agent such as phenazine methosulfate, phenazine ethosulfate, or menadione to assist and enhance formazan formation and assay sensitivity [22] . It is suggested that these electron intermediates shuttle into cells, pick up reducing equivalents from molecules such as nicotinamide adenine dinucleotide phosphate reduced (NADPH), and then shuttle back out into the culture medium where the tetrazolium is reduced to a soluble formazan.

HTS using various formulations of resazurin is becoming increasingly more common as a means of determining viability for drug screening. As a single reagent addition directly to cultures, it is readily adaptable to miniaturization in a robotic platform. Before use as a mammalian cell culture reagent for estimating viability, various protocols incorpo-rating either resazurin or methylthioninium chloride were developed during the 1930s as a simple and inexpensive means to determine bacterial content and grade raw milk supplies [26] . The original patented commercial product, commonly known as Alamar Blue, in addition to containing the redox dye resazurin, is thought to contain a mixture of stabilizer salts or poising agents. Their purpose is to maintain the redox potential of the culture medium alone in the range of 0.3 – 0.45 volts, thus keeping spontaneous background reduction in the absence of cells to a minimum. Alamar Blue is also believed to incorporate a second poising agent selected to minimize further cellular reduction of the fluorescent product (resorufin) to the colorless dihydroresorufin [27] .

Viability assays using resazurin are based on reduction of the oxidized blue dye, which has little intrinsic fluorescence, to its pink fluorescent resorufin product by living cells. At the same time, the visible light absorbance properties of the dye undergo a minor blue shift which can allow the assay to be monitored by absorbance, albeit with a loss of sensitivity due to the spectral overlap of the reduced and non-reduced dye. Cellular reduction of resazurin is believed to be accomplished by a pool of reductase or diaphorase-type enzymes derived from mitochondria and the cytosol [28] , many of which are shown to reduce MTT as well [18] . Confocal microscope investigation has demonstrated fluorescence that

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temporarily localizes primarily in the cytoplasm but can diffuse out and accumulate in the surrounding culture medium. The authors, however, could not negate reduction which initially may have taken place outside the cell, with resorufin diffusing into the cytoplasm and nucleus [28] . The Alamar Blue assay is marketed as non-toxic to cell cultures and is less likely to affect normal metabolism by not interfering with the electron transport chain [29] , thus allowing continuous monitoring for an extended period of time; however, this has been shown not to be true with all cell models [30] . For the purposes of HTS, however, the reagent is typically used as an end point reagent, in contact with the culture for a limited period of time (1 – 3 h) at the end of a drug treatment.

There is good correlation between results obtained with resazurin and other metabolism assays such as MTT; however, caution must be exercised when interpreting some results, particularly with reducing agents or chemicals that uncouple the mitochondrial electron transport system. Results from one screen using HepG2 cells and 117 compounds demonstrated that Alamar Blue compared favorably with MTT; both assays had nearly identical Z ′ factors [31] . This study showed that dicumarol, an anticoagulant and inhibitor of quinine reductase type 1, inhibited the fluorescence generated by Alamar Blue but enhanced the MTT absorbance signal with no obvious signs of toxicity as judged by micro-scopic observation using the Calcein AM viability test. In the same screen, MTT failed to detect daunorubicin toxicity, and the two assays were in significant discordance at estimating percentage viabilities with single dose trifluoperazine, thioridazine, and ellipticine treatments. Dicumarol enhance-ment of MTT signal, in a transient and exposure-dependent fashion, has been confirmed by others as well [32,33] . Others have observed increased MTT reduction with genistein treat-ments, which in parallel arrested cells in the G2/M cell cycle phase and caused a marked increase in cell volume and mitochondrial function [34] . In another comparison of resazurin to MTT, following treatment of three different brain cell lines with various highly purified advanced glycation end products, only the resazurin assay demonstrated cyotoxicity across all cell lines and treatments, with the MTT results varying between cell lines [35] . To confirm their findings, the authors relied upon a second assay, lactate dehydrogenase (LDH) release, to confirm toxicity across all cell lines and treatments. Ultimately many parameters influenced by experimental design, such as pH of the medium, glucose and NADPH content, MTT concen tration, and cell density, can impact formazan production and measurement, and should be addressed during assay validation experiments [36,37] .

2.2 Bioluminescent ATP assays Eukaryotic cells growing in vitro contain a relatively constant amount of ATP that is closely regulated to maintain homeostasis. During the process of cell death, there is a loss of ability to synthesize ATP and endogenous cytoplasmic

ATPases rapidly remove any remaining ATP. Measuring the amount of ATP from samples of cells in culture has been widely accepted as a valid marker of the number of viable cells present under most experimental conditions [38,39] . However, it should be recognized that change in ATP levels has also been used to measure properties such as activation of resting lymphocytes [40] .

The method of choice for measuring ATP is based on the ability of firefly luciferase to generate a luminescent signal. The original methods for measuring ATP required an acid extraction step to degrade endogenous ATPases and stabilize the amount of ATP present, followed by neutralizing the sample pH and subsequent combining with luciferin and native luciferase prepared from firefly ( Photinus pyralis ) abdomen. The resulting signal from this reaction was a flash of light lasting only a few seconds. Although this provided a sensitive manual method of detecting bacterial biomass or the number of viable eukaryotic cells in culture, sample manipulation and the short-lived signal limited the usefulness of this approach for measuring large numbers of samples. The development of luminometers with reagent injectors represented advancement but the number of samples that could be processed conveniently was still limited.

Two significant breakthroughs that enabled the broad acceptance of using the ATP assay as a marker for viable cells in the drug discovery process include: optimizing the reagent chemistry to create a glow signal and development of a stable form of luciferase. Optimizing the ingredients in luciferase assay reagents, to carefully balance the concen-trations of substrate, enzyme, and chemical inhibitors, resulted in the ability to control the rate of the luciferase reaction such that it produced a signal that would glow for several hours [41,42] . This achievement removed unwanted time restrictions and provided the convenience required for measuring large numbers of samples.

The second main improvement in the ATP assay technology was the engineering of a highly stable form of luciferase. Native firefly luciferase has limited stability. Environmental factors such pH and detergents can affect luciferase activity and limit reagent conditions desirable to develop robust homogeneous ATP assay reagents. To overcome this problem, Hall et al. used directed evolution techniques to develop mutant forms of luciferase from a related firefly, Photuris pennsylvanica , which were stable to environmental extremes and enabled development of a robust, homogeneous reagent for measuring ATP directly from cultured cells [43] .

The stable luciferase mutant was necessary for the develop-ment of a reagent that contains a sufficient concentration of detergent to immediately lyse cells upon addition, and ATPase inhibitor to stabilize the ATP released into the culture medium. The use of recombinant luciferase also allowed the production of large batches of enzyme in a controlled manufacturing environment resulting in a more consistent reagent batch-to-batch.

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The improved assay procedure was reduced to a single step to add the ATP detection reagent directly to cells in culture. After mixing and a brief 10 min equilibration period, the luminescent signal glows with a half-life generally greater than 5 h which provides convenience and flexibility for recording data from large batches of multi-well plates.

The luminescent ATP detection assay has several advantages and has become the method of choice for measuring cell viability in HTS labs [44-48] . The principal advantages of this method include the simplicity of the add-mix-measure homogeneous method, the speed of performing the assay, and the low interference from test compounds that typically interfere with fluorescent assay methods. In addition, it is the most sensitive microplate assay available for detecting viable cells in culture. The ATP assay can typically detect fewer than 10 cells per well, primarily because of the extremely low background luminescence present in biological samples. The reagent has adequate ambient temperature stability for use in an automated robotic platform and the assay is directly scalable from 96 to 384 to 1536 well formats.

Another main advantage of the ATP assay is that it immediately lyses cells on addition of the reagent, which provides information on the status of ATP at the instant the reagent is added. This is an advantage compared with tetrazolium reduction or resazurin reduction assays that can require hours of incubation with viable cells to convert the substrate to a colored or fluorescent product.

The disadvantages of this assay method include that it kills the cells and so the sample generally cannot be used for other purposes after treatment with the ATP detection reagent. Although there are several examples of multiplexing other assay methods, a sequential format is required. For example, measuring a luminescent Renilla genetic reporter assay first, followed by measuring ATP as a normalization control to measure viable cell number [49] . Another disadvantage of the ATP assay is variability resulting from temperature fluctuations. The enzymatic rate of luciferase and thus the luminescent signal is affected by temperature, so it is important to have a constant temperature environment to achieve the greatest reproducibility.

A potential disadvantage of all luciferase-based assays is the possibility that test compounds may inhibit luciferase, in this case causing a reduction in the signal without affecting cell viability or ATP content. Luciferase inhibitors in chemical libraries have been documented; however, their frequency in compound libraries is low [50] . The stabilized form of luciferase is used at high concentrations and the reagent formulations are designed to reduce compound interference.

2.3 Enzyme release-based cytotoxicity assays One of the most definitive methods for assessing cell death is to measure the leakage of cellular components from compromised cells into the culture medium. This is typically

achieved by measuring constitutive, conserved, and stable enzymatic activities released from dead cells. These assays are well suited for drug discovery efforts because the presence of biomarker activity is typically proof-positive for the presence of cytotoxicity and the assays are extraordinarily sensitive and amenable to high-density HTS environments.

LDH has long been favored as a marker of cell death for in vitro models [51,52] . LDH activity can be driven and indirectly measured by subjecting the sample to a coupled enzymatic chemistry reagent containing lactate, NAD + , diaphorase, and an appropriate redox dye such as resazurin, which produces either a change in absorbance or a shift in the fluorescence profile. Assay protocols have been historically limited in HTS, however, owing to the required removal of a small, cell-free aliquot of culture medium for testing in a separate plate. A significantly improved, truly homogeneous assay has been recently described which allows same-well measurement of LDH without the need for sampling [53] . This greatly simplified assay is now routinely implemented in high-density plate HTS screens [54,55] . The main detraction for this assay is that performance can be negatively impacted by serum supplemented medium. Therefore, cell-free, medium only controls must be included for meaningful analysis [56-58] . As with all activity assays, signal interferences exist, either by signal quenching, test compound auto-fluorescence, or by direct inhibition of the LDH assay chemistry [59-61] .

Highly sensitive bioluminescent cytotoxicity assays have been described which direct ATP generation through the activities of released ATP cycling enzymes. For instance, adenylate kinase (AK) activity can be measured in culture medium by providing ADP as a substrate to produce ATP [62,63] . Similarly, glyceraldehyde-3-phosphate dehydro-genase (GAPDH) activity can also be measured by providing a reaction cocktail containing coupled glycolytic pathway enzymes necessary for generating ATP [64,65] . Because these assays use a luminescent readout, they produce impressive signal-to-noise ratios in short time periods from relatively few numbers of dead cells. Although initially sensitive and linear, they do suffer from net signal decay as a function of limiting reactants and luciferase inactivation. This leads to loss of linearity across the cytotoxic spectrum, complicating the interpretation of the data beyond the recommended measurement times (15 min or less) [66] . In addition to the detriments related to the luciferase reaction chemistry, the suitability and validity of GAPDH as a cytotoxicity bio-marker is questionable in light of reports of its positive and negative modulation in the absence of membrane integrity disruptions [67-71] . One could similarly predict assay inter-ference with the AK chemistry by examining new chemical entities designed and directed at oncology-centric kinase inhibition [72-75] . Lastly, when compared with other biomarkers such as LDH, AK and GAPDH have relatively poor activity half-lives outside the cellular compartment [76] . Therefore, in extended experimental

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incubation time frames (greater than 24 h), actual cytotoxicity may be underestimated using these measures because of biomarker activity decay ( Figure 3 ).

A new cytotoxicity assay method was recently described which measures a distinct proteolytic profile produced by damaged cells [77] . In this method, a specific fluorogenic peptide is introduced directly into the culture medium where it is cleaved to generate a fluorescent product as a result of dead-cell proteolytic activity. Because the substrate is cell-impermeant, viable cells do not contribute to the fluorescence signal. This proteolytic assay method represents significant utility for drug discovery for a variety of reasons. First, more efficient, non-color quenching fluorogenic labels, such as rhodamine 110, can be used [78,79] . This allows same-well multiplexing with any other spectrally distinct assay chemistry, including a cell-permeable fluorogenic peptide substrate which measures the viable cells in an assay well [77] . Second, the reagent can be concentrated and delivered to assay wells in smaller volumes to accommodate more restricted-volume multiplex assay chemistries, such as optimized reagents for ATP or caspase detection [80-83] . This allows for elucidation of the mechanism for cytotoxicity and increases per-well informational content ( Figure 4 ). Last, the assay method has also been configured in a bioluminescent format using a peptide–luciferin conjugate and luciferase reaction components [66] . The glow-type luminescence signal generated from dead-cell protease activity is both stable and proportional to the number of dead cells for several hours (3 – 5 h). Therefore, both viable and non-viable cells can be measured in a sequential addition method whereby the cytotoxicity is measured first and the total cytotoxicity is measured after the introduction of a lysis reagent [82] . Because the luminescent signal derived from the experimental dead and detergent lysed pool is additive, the experimentally

viable population can be determined by subtracting the dead-cell signal values from the total signal [82] . The three multiplexed detection formats allowed by this technology (fluorescent/fluorescent, fluorescent/luminescent, or luminescent/luminescent) are all subject to different degrees of specific assay interferences from color quenching compounds, protease or luciferase inhibitors, or autofluore-scence. However, because of the ratiometric nature (inverse proportionality) of same-well viability and cytotoxicity measures, this technology has the ability to ‘flag’ assay inter-ferences for further study [76] . The ratiometric feature has additional benefits with regard to cell clumping or pipetting errors that can introduce significant variability and bias inherent in single point data sets with single parameter assays.

3. Conclusions

Although many assay options are available for assessment of cytotoxicity in vitro , relatively few are practical and have sufficient sensitivity, scalability, and robustness to be useful for drug screening or characterization efforts. For example, the neutral red viability assay is a classic example of a scientifically useful and validated method which is impractical and rarely used [84] in high-density formats owing to multiple additions, washes, and the requisite dye extraction step [44] . Of those with HTS utility, considerable practical differences exist with regard to their use. For example, assays measuring metabolic reductase activities have been used with great success and continue to be a useful and cost-effective staple of the industry. These reductase-based viability assays are well characterized, stable, cost-effective, and simple to use. The tetrazolium and resazurin reagents do require extended incubation (up to 4 h), however, to achieve measurable signals. Therefore, the final absorbance or

120

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00 2 4 6

Hours after cytotoxic event8 10 12

GAPDH Protease

LDHAK

Figure 3 . Cytotoxicity biomarkers based on enzymatic release have fi nite activity half-lives in culture medium. Biomarkers with poor stability may underestimate cytotoxicity during extended exposure periods.

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fluorescence measure represents an average of the viability over this incubation time frame. As well densities increase to 1536 and beyond, these assays also become less useful because of limitations inherent in the reagent chemistry [85,86] . Bioluminescent ATP viability assays offer advantages in high density formats owing to their high intrinsic sensitivity, low background values, impressive signal windows, and short incubation times (less than 10 min). Although there are considerably higher reagent costs associated with implementation of ATP assays, they provide rapid and reproducible results relative to the number of viable cells in an assay well [38,87,88] . ATP levels can fluctuate within cells

independent from cell death during various treatments [89,90] , and this may lead to under- or overestimation of the viable cell number. In addition, ATP assay chemistries must liberate the ATP from viable cells by a detergent. This means that ATP assays are a terminal end point in studies on cells after treatment because the lysis component is detrimental to most other assay chemistries and complicates other measure-ments. Both sets of viability assays do require a leap of inference, however, because they do not directly report the presence of cytotoxicity within an assay well. Viability assays merely report the number of viable cells remaining after treatment, and therefore may suggest cytotoxicity in the case

B. Viability EC50 = 6.89 μM

Cytotoxicity EC50 = 6.87 μM

Caspase EC50 = ND

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Cytoxicity EC50 = 1.9 nM

Viability EC50 = 1.9 nM

Caspase EC50 = 1.7 nM

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Figure 4 . Multi-parametric measures allow for elucidation of cytotoxic mechanism. Proteolytic biomarkers for viability and cytotoxicity were measured using the MultiTox-Fluor Assay. Caspase-3/7 activity was then measured in the same assay well using Caspase-Glo 3/7 Reagent (Promega Corporation). A. Paclitaxel treatment for 24 h resulted in a dose-dependent decrease in viability, increase in cytotoxicity, and an increase in caspase-3/7 activity consistent with apoptosis. B. Ionomycin treatment for 6 h resulted in a dose-dependent decrease in viability, increase in cytotoxicity, with no caspase-3/7 activation, which is consistent with primary necrosis.

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of cell-cycle arrest. Assay chemistries designed to detect enzymatic activities in the culture medium as a result of dying or dead cells are therefore proof-positive for a cytotoxic event. These assays are readily miniaturized into 384 and 1536 well formats, require only short incubation times (5 – 30 min), and are available in both fluorescent and luminescent signal formats. Ultimately, however, the practical utility of enzymatic activity assays relies on the kinetics of cell death in the model system, stability of the reporting signal, and the rate of degradation of the enzymatic biomarker (half-life) in the culture medium. Thus, these assays are useful within a defined window of time proximal to the cytotoxic event but may underestimate early stage cytotoxicity without membrane integrity changes. Conversely, during long exposure periods where early stage

death occurred and biomarker activity has waned, these assays may underestimate cytotoxicity levels. To mitigate the inherent liabilities of viability and cytotoxicity assays, it is considered good experimental practice to use either parallel or multiplexed assays to measure both parameters ( Figure 5 ). Although not always experimentally practical, and greatly dependent on the degree of statistical rigor required for the data sets generated, these complementary measures help to provide a complete understanding of the potential toxicity by preventing faulty interpretation of the data.

4. Expert opinion

No assay technology for detecting cytotoxicity in vitro is perfect. Strong arguments can be made for and against using

A.

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ATP EC50 = 63 nm

Metabolism reductase EC50 = 46 nm

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LDH release EC50 = 66 nmB.

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Figure 5 . Parallel orthogonal and/or multiplexed cytotoxicity and viability measures can strengthen data sets and prevent faulty interpretation. K562 cells were treated for 24 h with epoxomicin and subjected to viability and cytotoxicity assay chemistries. A. Metabolism reductase (resazurin) and ATP activity (in parallel); B. LDH activity; and C. multiplexed viability, cytotoxicity, and apoptosis measures.

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Table 1 . Comparison of viability and cytotoxicity assay chemistries.

Viability assays Cytotoxicity assays

ATP Resazurin MTS Protease LDH AK GAPDH Protease

Incubation time 5 min 1 – 4 h 1 – 4 h 30 min 10 min 5 min 5 min 15 min

Ease-of-use (homogeneous) Yes Yes Yes Yes Yes Yes Yes Yes

Detection method Lum. Fluor. Color. Fluor. or lum. Fluor. Lum. Lum. Fluor. or lum.

Interferences Yes Yes Yes Yes Yes Yes Yes Yes

Reagent stability/signal stability

Yes/yes Yes/yes Yes/yes Yes/yes Yes/yes ‡ Yes/initially Yes § /initially Yes/yes

Sensitivity *

96 well 384 well 1536 well

50 10 10

390 50 50

800 200 NA

50/50 50/10 50/10

800 200 NA

100 100 NA

500 500 NA

50/50 10/10 10/10

Cost $$ $ $ $$ $$ $$ $$ $$

Multiparametric assay options

Yes ¶ Yes # No Yes Yes # Unknown Unknown Yes

* Sensitivity is dependent on cell type and other experimental conditions. Sensitivity is expressed as cells/well in culture medium with 10% FBS. ‡ If stop solution is added. § If aliquoted and stored properly. ¶ If used last in sequence. # With deviation from standard protocol. $: Inexpensive; $$: More expensive; AK: Adenylate kinase; Color.: Colorimetric; FBS: Fetal bovine serum; Fluor.: Fluorescence; GAPDH: Glyceraldehyde-3-phosphate dehydrogenase; LDH: Lactate dehydrogenase; Lum.: Luminescence; MTS: 5-(3-Carboxymethoxyphenyl)-2-(4,5-dimethylthiazoly)-3-(4-sulfophenyl)tetrazolium, inner salt; NA: Information not available.

all of the chemistries described in this review during the process of drug discovery ( Table 1 ). Ultimately, preference should be given to a particular method based on the goals and constraints (fiscal, manpower, and man hours) of the study, and the degree to which the researcher is willing to accept risk for false positive or negative results. With this in mind, results from any combination of viability and cytotoxicity assays are rarely unanimous [91-95] .

Absorbance and fluorescence-based assays continue to bring value to cytotoxicity research efforts but confounding color-quenching or fluorescence interferences should not be underestimated, particularly with regard to natural product or natural product-like libraries of compounds [96,97] . Therefore, a movement is well underway to replace traditional absorbance and fluorescence chemistries with more efficient and sensitive green- or red-shifted fluorophores.

Many discovery groups have moved entirely away from absorbance- or fluorescence-based methods to luminescent detection systems. The remarkable expansion in the use of bioluminescent assay chemistries for cytotoxicity testing is driven by a variety of factors. First, natural bioluminescence is confined to a handful of insect and sea-dwelling invertebrates, so mammalian systems are free from artifactual interferences resident in fluorescence assays owing to autofluorescence [98] . Second, luminescence is an inherently more sensitive detection format because photons are generated chemically rather than being produced by flooding

the assay well with a fluorescence excitation source [99] . Third, the use of luminescence as a reporting system has been thoroughly examined and validated now in a variety of drug screening environments and has been proven to be useful. Last, more luminescent chemistries have become commercially available in the last decade for the detection of biomarkers of cytotoxicity or eventual cytotoxicity. In addition to ATP, ATP-cycling, or protease release assays described previously in this review, there are now strongly predictive and sensitive cell-based bioluminescent assays for glutathione levels, caspases-3/7, -8, and -9 and cytochrome P450 activity that measure and may help elucidate specific pathways and/or pre-cytotoxicity mechanism-based mediators of global cell health [100] . Although luminescence typically offers a considerable improvement over colorimetric or fluorometric detection systems, it is also susceptible to interferences from color quenching compounds or by direct inhibition of luciferase [101] . Therefore, luciferase-based detection systems have the capacity to be negatively impacted by focused libraries containing ATP analogues. In practice, however, numerous examples exist for successful kinase inhibition screens using luciferase-based detection [50,102] .

Historically, assay-specific interferences resulting in false positive or negative hits have been mitigated by orthogonal assay chemistries conducted in parallel or secondary screens. This approach not only addresses statistical concerns and result validity but also introduces well-to-well variability and

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additional costs associated with duplication of effort. Therefore, a variety of multi-parametric and homogeneous, same-well methods have been developed which use varying degrees of reagent as well as assay chamber and detecting instrument sophistication. For example, the authors [103] and others [104] have focused on both conventional and innovative chemistries in compatible, multiplexed combinations. A significant advantage of this approach is that these plate-based chemistries provide spectrally distinct orthogonal or inversely proportional measures of cytotoxicity that can be measured using standard fluorometry or luminometry. The validity and biochemical principles inherent in these measures are familiar, accessible, accepted, and readily adapted to robotics platforms with little or no requirement for specialized training, equipment, or data analysis packages. Furthermore, the statistical probability of a particular compound interfering with multiple platforms of detection is therefore lessened by inherent internal control [105,106] . The use of this internal control is not only useful for examination of potential cytotoxicity but can also be extended to normalization of cell number, which may be of practical use in experiments studying genetic reporter responses or siRNA knockdown. Finally, global cytotoxicity measures can often be easily scaled and combined with mechanism-specific measures such as caspase activation.

Exciting advances in automated imaging and computer-aided data analysis have driven multi-parametric, ‘phenotypic’ cell-based assays into the forefront of drug discovery efforts [107,108] . High content screening (HCS) technology permits systematic analysis of spatial and quantitative changes in cellular morphology, genes, and proteins as a result of compound or treatment effects [109] . These measures are made possible by using nearly unlimited combinations of classical dyes and DNA intercalators with specific antibodies and fluorescently tagged proteins. Although numerous examples of successful cytotoxicity screens exist for HCS, the chief benefit of this specialized technology is that it not only permits active compound identification but also helps elucidate target

identification [110] . A principal challenge for widespread implementation of the technology has been the complexity of data analysis, the size of data files, and instrumentation acquisition costs. Although the HCS technology is relatively immature with respect to conventional cytotoxicity and viability chemistries, it promises to significantly impact the future of drug discovery efforts.

In conclusion, considerable interest is now given to label-free technologies because they eliminate concerns related to fluorescence and luminescence interferences, and do not require parameter-specific, exogenously added probes [111-113] . Several systems have been developed using cellular dielectric spectroscopy (CDS) where physiological events are monitored through complex measurement of impedance changes within specialized assay wells [113,114] . Events that affect morphological or cellular adherence, such as early or late-stage cytotoxicity, can be identified by their influence on the magnitude of electrical conductivity. Advanced adaptations, such the system developed by Bionas, use silicon chips equipped with parametric sensors, and feature the ability to make dynamic and continuous measurements of three key metabolic parameters (oxygen consumption, extracellular acidification rate, and adhesion properties) for up to several days. These active, online measurements of metabolic parameters are particularly attractive because they obviate limitations associated with end point assays which have less tolerance for transient and temporal effects [115] . Therefore, real-time kinetic measure-ment of cytotoxicity represents a principal advantage over fluorescent or luminescent systems. Unfortunately, at present these systems offer only low to modest throughputs (6 – 96 wells at a time), have unproven cost-effectiveness, and will require further development for miniaturization and widespread implementation. Nevertheless, label-free technologies represent a great hope for the future.

Declaration of interest

The authors are employees of Promega Corporation.

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57. Hasinoff B, Schroeder P, Patel D. The metabolites of the cardioprotective drug dexrazoxane do not protect myocytes from doxorubicin-induced cytotoxicity. Mol Pharmacol 2003 ; 64 : 67

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• Another early paper describing cytotoxicity biomarkers.

63. Squirrel D, Murphy J. Rapid detection of very low number of micro-organisms using adenylate kinase as a cell marker.

A practical guide to industrial uses of ATP luminescence in rapid microbiology. 1997 : 107 -13

64. Corey MJ, et al. A very sensitive coupled luminescent assay for cytotoxicity and complement-mediated lysis. J Immunol Methods 1997 ; 207 : 43 -51

65. Ogbomo H, Hahn A, Geiler J, et al. NK sensitivity of neuroblastoma cells determined by a highly sensitive coupled luminescent method. Biochem Biophys Res Commun 2006 ; 339 : 375 -9

66. Cho M-H, Niles A, Huang R, et al. A bioluminescent cytotoxicity assay for assessment of membrane integrity using a proteolytic biomarker. Toxicol In Vitro 2008 ; 22 : 1099 -106

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68. Du Z-X, Wang H-Q, Zhang H-Y, et al. Involvement of glyceraldehyde-3-phosphate dehydrogenase in tumor necrosis factor-related apoptosis-inducing ligand mediated death of thyroid cancer cells. Endocrinology 2007 ; 148 : 4352 -61

69. Dimmeler S, Ankarcrona M, Nicotera P, et al. Exogenous nitric oxide (NO) generation or IL-1 β -induced intracellular NO production stimulates inhibitory auto-ADP-ribosylation of glyceraldehyde-3-phosphate dehydrogenase in RINm5F cells. J Immunol 1993 ; 150 : 2964 -71

70. Ito Y, Pagano K, Tornheim P, et al. Oxidative stress increases glyceraldehyde-3-phosphate dehydrogenase mRNA levels in isolated rabbit aorta. Am J Physiol Heart Circ Physiol 1996 ; 270 : H81 -7

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72. Carvajal R, Tse A, Schwartz G. Aurora kinases: new targets for cancer therapy. Clin Cancer Res 2006 ; 12 : 6869 -75

73. Heinrich M, Blanke C, Druker B, et al. Inhibition of KIT tyrosine kinase activity: a novel molecular approach to the treatment of KIT-positive malignancies. J Clin Oncol 2002 ; 20 : 1692 -703

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74. Schöffski P, Dumez H, Clement A, et al. Emerging role of tyrosine kinase inhibitors in the treatment of advanced renal cell cancer: a review. Ann Oncol 2006 ; 17 : 1185 -96

75. Schmit T, Nihal A. Regulation of mitosis via mitotic kinases: new opportunities for cancer management. Mol Cancer Ther 2007 ; 6 : 1920 -31

• Excellent perspective on targeted anti-cancer strategies.

76. Niles A, Scurria M, Bernad L, et al. Measure relative numbers of live and dead cells and normalize assay data to cell number. Cell Notes 2007 ; 18 : 15 -20

77. Niles A, Moravec R, Hesselbeth P, et al. A homogeneous assay to measure live and dead cells in the same sample by detecting different protease markers. Anal Biochem 2007 ; 366 : 197 -206

•• This paper describes the discovery, characterization, and the multiplexed utility of new proteolytic biomarkers.

78. Liu J, Bhalgat M, Zhang C, et al. Fluorescent molecular probes V: a sensitive caspase-3 substrate for fl uorometric assays. Bioorg Med Chem Lett 1999 ; 9 : 3231 -6

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80. Niles A, Moravec R, Scurria M, et al. MultiTox-fl uor multiplex cytotoxicity assay technology. Cell Notes 2006 ; 15 : 11 -5

81. Niles A, Worzela T, Scurria M, et al. Multiplexed viability, cytotoxicity and apoptosis assays for cell-based screening. Cell Notes 2006 ; 16 : 12 -5

• An easy reading description of multiplexed measures in screening.

82. Niles A, Scurria M, Bernad L, et al. Using protease biomarkers to measure viability and cytotoxicity. Cell Notes 2007 ; 19 : 16 -20

83. Worzella T, Busch M, Niles A. High throughput automation of multiplexed cell viability assays for viability and cytotoxicity. Cell Notes 2008 ; 20 : 26 -9

84. Jones P, King A. High throughput screening (HTS) for phototoxicity hazard using the in vitro 3T3 neutral red uptake assay. Toxicol In Vitro 2003 ; 17 : 703 -8

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• A good description of the challenges inherent in HTS scaling.

87. Maehara Y, Anai H, Tamada R, et al. The ATP assay is more sensitive than the succinate dehydrogenase inhibition test for predicting cell viability. Eur J Cancer Clin Oncol 1987 ; 23 : 273 -6

88. Slater K. Cytotoxicity tests for high-throughput drug discovery. Cur Opin Biotechnol 2001 ; 12 : 70 -4

89. Sanchez-Alcazar J, Ruiz-Cabello J, Hernandez-Munoz I, et al. Tumor necrosis factor- α increases ATP content in metabolically inhibited L929 cells preceding cell death. J Biol Chem 1997 ; 272 : 30167 -77

90. Shchepina L, Pletjushhkina O, Averisyan A, et al. Oligomycin, inhibitor of the F o part of H + -ATP synthase, suppresses the TNF-induced apoptosis. Oncogene 2002 ; 21 : 8149 -57

91. Jondeau A, Dahbi L, Bani-Estivals M-H, et al. Evaluation of the sensitivity of three sublethal cytotoxicity assays in human HepG2 cell line using water contaminants. Toxicology 2006 ; 226 : 218 -28

92. Weyermann J, Lochmann D, Zimmer A. A practical note on the use of cytotoxicity assays. Int J Pharm 2005 ; 288 : 369 -76

• A comparison of four different detection chemistries with four cytotoxic mechanisms.

93. Puttonen K, Lehtonen S, Lampela P, et al. Different viabilities and toxicity types after 6-OHDA and Ara-C exposure evaluated by four assays in fi ve cell lines. Toxicol In Vitro 2008 ; 22 : 182 -9

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95. Ulukaya E, Ozdikicioglu F, Yilmaztepe-Oral A, et al. The MTT assay yields a relatively lower result of growth inhibition than the ATP assay depending on the chemotherapeutic drugs tested. Toxicol In Vitro 2008 ; 22 : 232 -9

96. Ausseil F, Samson A, Aussagues Y, et al. High-throughput bioluminescence screening of ubiquitin-proteasome pathway

inhibitors from chemical and natural sources. J Biomol Screen 2007 ; 12 : 106 -16

97. Turek-Etienne T, Lei M, Terracciano J, et al. Use of red-shifted dyes in a fl uorescence polarization AKT kinase assay for detection of biological activity in natural product extracts. J Biomol Screen 2004 ; 9 : 52 -61

98. O’Brien M, Daily W, Hesselberth E, et al. Homogeneous, bioluminescent protease assays: caspase-3 as a model. J Biomol Screen 2005 ; 10 : 137 -48

• A concise description of modifi ed luciferin-based, caspase detection chemistries.

99. Fan F, Wood K. Bioluminescent assays for high-throughput screening. Assay Drug Dev Technol 2007 ; 5 : 127 -36

•• This review presents the arguments for implementing luminescence detection chemistries in HTS.

100. Cali J, Niles A, Valley M, et al. Bioluminescent assays for ADMET. Expert Opin Drug Metab Toxicol 2008 ; 4 : 103 -20

•• A strong survey of available bioluminescent methods for ADMET.

101. Pratt S, David C, Black-Schaefer C, et al. A strategy for discovery of novel broad-spectrum antibacterials using a high-throughput Streptococcus pneumoniae transcription/translation screen. J Biomol Screen 2004 ; 9 : 3 -11

102. Schröter T, Minond D, Scampavia L, et al. Comparison of miniaturized time-resolved fl uorescence resonance energy transfer and enzyme-coupled luciferase high-throughput screening assays to discover inhibitors of Rho-kinase II (ROCK-II). J Biomol Screen 2008 ; 13 : 17 -28

103. Niles A, Moravec R, Riss T. Multiplex caspase activity and cytotoxicity assays. Methods Mol Biol 2008 ; 414 : 151 -62

•• Methods are provided for combining viability, cytotoxicity, and caspase detection chemistries.

104. Hallis T, Kopp A, Gibson J, et al. An improved β -lactamase reporter assay: multiplexing with a cytotoxicity readout for enhanced accuracy of hit identifi cation. J Biomol Screen 2007 ; 12 : 635 -44

• This paper demonstrates multiplex normalization of genetic reporter data.

105. Grant S, Sklar J, Cummings R. Development of novel assays for proteolytic enzymes using rhodamine-based

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fl uorogenic substrates. J Biomol Screen 2002 ; 7 : 531 -40

106. Nieuwenhuijsen B, Huang Y, Wang Y, et al. A dual luciferase multiplexed high-throughput screening platform for protein-protein interactions. J Biomol Screen 2003 ; 8 : 676 -84

107. Giuliano K, Haskins J, Taylor D. Advances in high content screening for drug discovery. Assay Drug Dev Technol 2003 ; 1 : 565 -77

108. Abraham V, Taylor D, Haskins J. High content screening applied to large-scale biology. Trends Biotechnol 2004 ; 22 : 15 -22

109. Giuliano K, DeBiasio R, Dunlay R, et al. High-content screening: a new approach to easing key bottlenecks in the drug discovery process. J Biomol Screen 1997 ; 2 : 249 -59

110. Wilson C, Si Y, Thompson C, et al. Identifi cation of a small molecule that induces mitotic arrest using a simplifi ed high-content screening assay and data analysis method. J Biomol Screen 2006 ; 11 : 21 -8

111. Ciambrone G, Liu V, Lin D, et al. Cellular dielectric spectroscopy: a powerful new approach to label-free cellular analysis. J Biomol Screen 2004 ; 9 : 467 -80

112. Zhu J, Wang X, Xu X, et al. Dynamic and label-free monitoring of natural killer cell cytotoxicity using electronic cell sensor arrays. J Immunol Methods 2006 ; 309 : 25 -33

113. Xing J, Zhu L, Gabos S, et al. Microelectric cell sensor assay for detection of cytotoxicity and prediction of acute toxicity. Toxicol In Vitro 2006 ; 20 : 995 -1004

114. Peters M, Knappenberger K, Wilkens D, et al. Evaluation of cellular dielectric spectroscopy, a whole-cell, label-free technology for drug discovery on G i -coupled GPCRs. J Biol Screen 2007 ; 12 : 312 -9

115. Thedinga E, Kob A, Holst H, et al. Online monitoring of cell metabolism for studying pharmacodynamic effects. Toxicol Appl Pharm 2007 ; 220 : 33 -44

• A view of multi-parametric assays of the future.

Affi liation Andrew L Niles † 1 MS , Richard A Moravec 2 BS & Terry L Riss 3 PhD † Author for correspondence 1 Senior Research Scientist Promega Corporation, Research and Development, 2800 Woods Hollow Road, Madison, Wisconsin, 53711, USA Tel: +1 608 247 4330, ext. 1447 ; Fax: +1 608 298 4818 ; E-mail: [email protected] 2 Senior Research Scientist Promega Corporation, Research and Development, 2800 Woods Hollow Road, Madison, Wisconsin, 53711, USA 3 Director of Project Management Promega Corporation, Research and Development, 2800 Woods Hollow Road, Madison, Wisconsin, 53711, USA

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