A live-cell assay for the real-time assessment of extracellular ATP levels.

Extracellular ATP (eATP) is a potent damage associated molecular pattern (DAMP) molecule known to exert profound effects on the innate and adaptive immune responses. As such, it has become an important biomarker for studying means to pro-actively modulate inflammatory processes. Unfortunately, traditional methodologies employed for measuring eATP require cumbersome supernatant sampling, onerous time courses, or unnecessary duplication of effort. Here we describe a new reagent that is tolerable to test cells in extended exposures and enables a fully homogeneous assay method for real-time determinations of extracellular ATP levels. The reagent is introduced into assay plates containing cells at the time of stimulus introduction. The real-time feature of the format allows for sensitive, continuous accounting of eATP levels in the test model over at least 24 h. This work details our efforts to create and characterize this new reagent and to validate utility by demonstrating its use with multiple cell lines and chemically diverse eATP induction stimuli.

A real-time, bioluminescent annexin V assay for the assessment of apoptosis.

Apoptosis is an important and necessary cell death program which promotes homeostasis and organismal survival. When dysregulated, however, it can lead to a myriad of pathologies from neurodegenerative diseases to cancer. Apoptosis is therefore the subject of intense study aimed at dissecting its pathways and molecular mechanisms. Although many assay methods exist for confirming whether an apoptotic response has occurred in vitro, most methods are destructive and involve laborious operator effort or specialized instrumentation. Here we describe a real-time, no-wash, microplate method which utilizes recombinant annexin V fusion proteins containing evolved binary subunits of NanoBiT™ luciferase. The fusion proteins, a time-released enzymatic substrate, a necrosis detection dye and exogenous calcium ions are delivered via an optimized and physiologically inert reagent directly to cells in culture at the time of treatment or dosing. Luminescent signals proportional to phosphatidylserine (PS) exposure and fluorescent signals generated as a result of loss of membrane integrity are then collected using a standard multimode plate reader at scheduled intervals over the exposure. The resulting luminescent and fluorescent data are then used to define the kinetics and magnitude of an apoptotic response. This study details our efforts to develop, characterize, and demonstrate the features of the assay by providing relevant examples from diverse cell models for programmed cell death.

A Nondestructive, Real-Time Annexin V Apoptosis Assay.

This chapter describes a simple, nondestructive, annexin V apoptosis detection method that can be employed in real time over a 48-h test exposure. The real-time functionality allows for temporal resolution of apoptotic and cell death responses during the test exposure and obviates the need for onerous sample preparation and time course protocols associated with other annexin V methods. Further, this technique is eminently accessible to a wide range of laboratories because it does not require flow cytometry or other cytometric methods. It was developed for use with a variety of microplate well densities and with standard multimodal plate readers. The central feature of this assay is that it continuously reports the residency status of phosphatidylserine (PS) on the exofacial surface of a cell as it is translocated from the inner membrane leaflet during the apoptotic process. This homogenous, no-wash assay is made possible by two optimized and distinct annexin V fusion proteins which contain complementing NanoBiT™ luciferase enzyme subunits, a time-released luciferase substrate, and a fluorescent membrane integrity reagent. During an apoptotic event, the luminescent signal arising from an assay well is proportional to the number of cells with PS exposure, and fluorescence intensity correlates with the degree of cell death (secondary necrosis). Conversely, untreated cells contribute negligible luminescent or fluorescent signals throughout the time course. The data collected from these assay measures provide for both standard potency determinations and kinetic characterization of dose- and agent-dependent apoptotic responses, from early through late phases.

A Real-Time, Bioluminescent Apoptosis Assay.

This chapter describes a real-time, bioluminescent apoptosis assay technique, which circumvents the well-documented "timing condundrum" encountered when employing traditional apoptosis detection chemistries after exposures with inducers of unknown potential. The assay continuously reports the translocation of phosphatidylserine (PS) from the inner membrane leaflet of a cell to the exofacial surface during apoptosis. This homogenous, no-wash, plate-based assay is made possible by two different annexin V fusion proteins, which contain complementing NanoBiT™ luciferase enzyme subunits, a time-released luciferase substrate, and a fluorescent membrane integrity reagent. During apoptosis, luminescence signal is proportional to PS exposure and fluorescence intensity correlated with the degree of secondary necrosis. Altogether, the measures provide exquisite kinetic resolution of dose- and agent-dependent apoptotic responses, from early through late phases. At exposure termination, other compatible reagents can be applied to measure additional orthogonal correlates of cell health.

Treating Cells as Reagents to Design Reproducible Assays.

The reproducibility of high-throughput cell-based assays is dependent on having a consistent source of cells for each experiment. Developing an understanding of the nature of cells growing in vitro and factors that influence their responsiveness to test compounds will contribute to the development of reproducible cell-based assays. Using good cell culture practices and establishing standard operating procedures (SOPs) for handling cultures can eliminate several potential contributors to variability in the responsiveness and performance of cells. The SOPs for handling each cell type must have clear and detailed instructions that can be understood and followed among different laboratories. The SOPs should include documenting the source of cells and authenticating their identity, both of which have become required to achieve peer acceptance of experimental data. Variability caused by biological issues such as phenotypic drift can be reduced by using standardized subculture procedures or using cryopreserved cells to set up experiments. Variability caused by inconsistent dispensing of cells per well and edge effects can be identified by measuring how many cells are present and whether they are alive or dead. Multiplex methods for real-time measurement of viable or dead cell number in each sample can be used for normalizing data and determining if proliferation or cytotoxicity has occurred during the experiment. Following good cell culture practices will go a long way toward executing reproducible cell-based assays. Resources will be included describing good cell culture practices, cell line authentication, and multiplex determination of cell number as an internal control.

Caspase activity assays.

Caspase activity assays in multi-well plate formats represent powerful tools for understanding experimental modulation of the apoptotic response. These assays are configured to exploit functional, biochemical, and temporal differences in substrate specificity and selectivity, which are useful in defining the magnitude and mechanism of a compound or treatment effect. New advances in fluorescent and luminescent chemistries now enable single addition "add-mix- measure" determinations of caspase activity directly in the sample plate with unprecedented sensitivity. Unlike other more cumbersome or laborious techniques, caspase activity induction or inhibition measures are quantifiable and definitive. The highlighted techniques in this chapter are cost efficient and allow for the rapid exploration of thousands of combinations and conditions.

Multiplex caspase activity and cytotoxicity assays.

Multiplexed assay chemistries provide for multiple measurements of cellular parameters within a single assay well. This experimental practice not only is more cost efficient but provides more informational content about a compound or treatment. For instance, multiplexed caspase activity assays can help establish the kinetics and magnitude of initiator and effector caspase induction by candidate compounds or treatments. The ability to combine the activity profiles within the same sample provides a level of normalization not possible with parallel assays. Furthermore, multiplexing caspase activity assays with viability and/or cytotoxicity assays can support conclusions regarding cytotoxic mechanism and provide normalization that may help correct for differences in cell number.

A bioluminescent cytotoxicity assay for assessment of membrane integrity using a proteolytic biomarker.

Measurement of cell membrane integrity has been widely used to assess chemical cytotoxity. Several assays are available for determining cell membrane integrity including differential labeling techniques using neutral red and trypan blue dyes or fluorescent compounds such as propidium iodide. Other common methods for assessing cytotoxicity are enzymatic "release" assays which measure the extra-cellular activities of lactate dehydrogenase (LDH), adenylate kinase (AK), or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) in culture medium. However, all these assays suffer from several practical limitations, including multiple reagent additions, scalability, low sensitivity, poor linearity, or requisite washes and medium exchanges. We have developed a new cytotoxicity assay which measures the activity of released intracellular proteases as a result of cell membrane impairment. It allows for a homogenous, one-step addition assay with a luminescent readout. We have optimized and miniaturized this assay into a 1536-well format, and validated it by screening a library of known compounds from the National Toxicology Program (NTP) using HEK 293 and human renal mesangial cells by quantitative high-throughput screening (qHTS). Several known and novel membrane disrupters were identified from the library, which indicates that the assay is robust and suitable for large scale library screening. This cytotoxicity assay, combined with the qHTS platform, allowed us to quickly and efficiently evaluate compound toxicities related to cell membrane integrity.

Bioluminescent assays for ADMET.

Bioluminescent assays couple a limiting component of a luciferase-catalyzed photon-emitting reaction to a variable parameter of interest, while holding the other components constant or non-limiting. In this way light output varies with the parameter of interest. This review describes three bioluminescent assay types that use firefly luciferase to measure properties of drugs and other xenobiotics which affect their absorption, distribution, metabolism, elimination and toxicity. First, levels of the luciferase enzyme itself are measured in gene reporter assays that place a luciferase cDNA under the control of regulatory sequences from ADMET-related genes. This approach identifies activators of nuclear receptors that regulate expression of genes encoding drug-metabolizing enzymes and drug transporters. Second, drug effects on enzyme activities are monitored with luminogenic probe substrates that are inactive derivatives of the luciferase substrate luciferin. The enzymes of interest convert the substrates to free luciferin, which is detected in a second reaction with luciferase. This approach is used with the drug-metabolizing CYP and monoamine oxidase enzymes, apoptosis-associated caspase proteases, a marker protease for non-viable cells and with glutathione-S-transferase to measure glutathione levels in cell lysates. Third, ATP concentration is monitored as a marker of cell viability or cell death and as a way of identifying substrates for the ATP-dependent drug transporter, P-glycoprotein. Luciferase activity is measured in the presence of a sample that supplies the requisite luciferase substrate, ATP, so that light output varies with ATP concentration. The bioluminescent ADMET assays are rapid and sensitive, amenable to automated high-throughput applications and offer significant advantages over alternative methods.

"Multiplexed viability, cytotoxicity, and caspase activity assays".

Multiplexed assay chemistries provide for multiple measurements of cellular parameters within a single assay well. This experimental practice is not only more cost efficient, but also provides more information about a compound or treatment. The ability to combine the activity profiles within the same sample provides a level of normalization not possible with parallel assays. Furthermore, multiplexing caspase activity assays with viability and/or cytotoxicity assays can support conclusions regarding cytotoxic mechanism and provide normalization, which may help correct for differences in cell number.

Target engagement and drug residence time can be observed in living cells with BRET.

The therapeutic action of drugs is predicated on their physical engagement with cellular targets. Here we describe a broadly applicable method using bioluminescence resonance energy transfer (BRET) to reveal the binding characteristics of a drug with selected targets within intact cells. Cell-permeable fluorescent tracers are used in a competitive binding format to quantify drug engagement with the target proteins fused to Nanoluc luciferase. The approach enabled us to profile isozyme-specific engagement and binding kinetics for a panel of histone deacetylase (HDAC) inhibitors. Our analysis was directed particularly to the clinically approved prodrug FK228 (Istodax/Romidepsin) because of its unique and largely unexplained mechanism of sustained intracellular action. Analysis of the binding kinetics by BRET revealed remarkably long intracellular residence times for FK228 at HDAC1, explaining the protracted intracellular behaviour of this prodrug. Our results demonstrate a novel application of BRET for assessing target engagement within the complex milieu of the intracellular environment.

Discovering protein interactions and characterizing protein function using HaloTag technology.

Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous proteomes, including those from viruses, bacteria, and yeast. In comparison, analysis of the human proteome lags behind, partially due to the sheer number of proteins which must be studied, but also the complexity of networks and interactions these present. To specifically address the challenges of understanding the human proteome, we have developed HaloTag technology for protein isolation, particularly strong for isolation of multiprotein complexes and allowing more efficient capture of weak or transient interactions and/or proteins in low abundance. HaloTag is a genetically encoded protein fusion tag, designed for covalent, specific, and rapid immobilization or labelling of proteins with various ligands. Leveraging these properties, numerous applications for mammalian cells were developed to characterize protein function and here we present methodologies including: protein pull-downs used for discovery of novel interactions or functional assays, and cellular localization. We find significant advantages in the speed, specificity, and covalent capture of fusion proteins to surfaces for proteomic analysis as compared to other traditional non-covalent approaches. We demonstrate these and the broad utility of the technology using two important epigenetic proteins as examples, the human bromodomain protein BRD4, and histone deacetylase HDAC1. These examples demonstrate the power of this technology in enabling the discovery of novel interactions and characterizing cellular localization in eukaryotes, which will together further understanding of human functional proteomics.

Validation of a HTS-amenable assay to detect drug-induced mitochondrial toxicity in the absence and presence of cell death.

Drug-induced mitochondrial dysfunction is known to contribute to late stage compound attrition. Recently, assays that identify mitochondrial dysfunction have been developed but many require expensive reagents, specialized equipment, or specialized expertise such as isolation of mitochondria. Here, we validate a new 384-well format cell-based dual parameter assay that uses commonly available detection methods to measure both mitochondrial toxicity and cytotoxicity. In our initial evaluation, antimycin A, CCCP, nefazodone, flutamide, and digitonin were tested in K562 cells in both glucose- and galactose-supplemented media with a 2h incubation. The assay was able to correctly differentiate these compounds into mitochondrial toxicants and non-mitochondrial toxicants, and had excellent reproducibility. We next tested 74 compounds in K562 cells in both types of media and show that the assay was able to correctly identify some of the compounds as mitochondrial toxicants. Moreover, the assay could be simplified, without loss of information, by using K562 cells in galactose-containing medium alone. This simple, robust assay can be positioned as a rapid, early readout of mitochondrial and cellular toxicity. However, since the assay fails to identify some mitochondrial toxicants, further assays may be required to detect mitochondrial toxicity once lead compounds have been selected.

Cytotoxicity testing: measuring viable cells, dead cells, and detecting mechanism of cell death.

Testing the effects of compounds on the viability of cells grown in culture is widely used as a predictor of potential toxic effects in whole animals. Among the several alternative assays available, measuring the levels of ATP is the most sensitive, reliable, and convenient method for monitoring active cell metabolism. However, recently developed combinations of methods have made it possible to collect more information from in vitro cytotoxicity assays using standard fluorescence and luminescence plate readers. This chapter describes two assay methods. The first utilizes beetle luciferase for measuring the levels of ATP as a marker of viable cells. The second more recently developed multiplex method relies on selective measurement of three different protease activities as markers for viable, necrotic, and apoptotic cells. Data analysis from the measurement of three marker protease activities from the same sample provides a useful tool to help uncover the mechanism of cell death and can serve as an internal control to help identify assay artifacts.

A bioluminogenic HDAC activity assay: validation and screening.

Histone deacetylase (HDAC) enzymes modify the acetylation state of histones and other important proteins. Aberrant HDAC enzyme function has been implicated in many diseases, and the discovery and development of drugs targeting these enzymes is becoming increasingly important. In this article, the authors report the evaluation of homogeneous, single-addition, bioluminogenic HDAC enzyme activity assays that offer less assay interference by compounds in comparison to fluorescence-based formats. The authors assessed the key operational assay properties including sensitivity, scalability, reproducibility, signal stability, robustness (Z'), DMSO tolerance, and pharmacological response to standard inhibitors against HDAC-1, HDAC-3/NcoR2, HDAC-6, and SIRT-1 enzymes. These assays were successfully miniaturized to a 10 µL assay volume, and their suitability for high-throughput screening was tested in validation experiments using 640 drugs approved by the Food and Drug Administration and the Hypha Discovery MycoDiverse natural products library, which is a collection of 10 049 extracts and fractions from fermentations of higher fungi and contains compounds that are of low molecular weight and wide chemical diversity. Both of these screening campaigns confirmed that the bioluminogenic assay was high-throughput screening compatible and yielded acceptable performance in confirmation, counter, and compound/extract and fraction concentration-response assays.

In vitro viability and cytotoxicity testing and same-well multi-parametric combinations for high throughput screening.

In vitro cytotoxicity testing has become an integral aspect of drug discovery because it is a convenient, costeffective, and predictive means of characterizing the toxic potential of new chemical entities. The early and routine implementation of this testing is testament to its prognostic importance for humans. Although a plethora of assay chemistries and methods exist for 96-well formats, few are practical and sufficiently sensitive enough for application in high throughput screening (HTS). Here we briefly describe a handful of the currently most robust and validated HTS assays for accurate and efficient assessment of cytotoxic risk. We also provide guidance for successful HTS implementation and discuss unique merits and detractions inherent in each method. Lastly, we discuss the advantages of combining specific HTS compatible assays into multi-parametric, same-well formats.

Update on in vitro cytotoxicity assays for drug development.

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.

A homogeneous assay to measure live and dead cells in the same sample by detecting different protease markers.

A method to simultaneously determine the relative numbers of live and dead cells in culture by introducing a combination of two fluorogenic substrates or a fluorogenic and a luminogenic protease substrate into the sample is described. The method is based on detection of differential ubiquitous proteolytic activities associated with intact viable cells and cells that have lost membrane integrity. A cell-permeable peptide aminofluorocoumarin substrate detects protease activity restricted to intact viable cells. Upon cell death, the viable cell protease marker becomes inactive. An impermeable peptide rhodamine 110 (or aminoluciferin) conjugated substrate detects protease activity from nonviable cells that have lost membrane integrity. The multiplex assay can detect 200 dead cells in a population of 10,000 viable cells. The protease substrate reagents do not damage viable cells over the course of the assay, thus the method can be multiplexed further with other assays in a homogeneous format. Ratiometric measurement of viable and dead cells in the same sample provides an internal control that can be used to normalize data from other cell-based assays.