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Sittampalam GS, Coussens NP, Nelson H, et al., editors. Assay Guidance Manual [Internet]. Bethesda (MD): Eli Lilly & Company and the National Center for Advancing Translational Sciences; 2004-.

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Inhibition of Protein-Protein Interactions: Non-Cellular Assay Formats

, PhD, , PhD, , PhD, , and .

Author Information

, PhD,1,* , PhD,2,* , PhD,3 ,3 and 3.

1 University of California San Francisco, San Francisco, CA
2 Harvard NeuroDiscovery Center, Boston, MA
3 Emory University School of Medicine, Atlanta, GA

Published ; Last Update: October 1, 2012.


Protein-protein interactions (PPI) are critical in cellular signal transductions that play a key role in both normal and abnormal functions in cells. Therefore, modulating the activity of these interactions is a major focus in drug discovery research. In this chapter, the authors address the development, optimization and validation of HTS assays to identify small molecule modulators of PPI. They also discuss the sources of artifacts, detailed accounts of assay technologies compatible with HTS for PPI and validating the inhibition of PPI. An extensive set of references is provided, and is a must read for beginners and a review for experienced investigators.

Overview and Introduction


Protein-protein interactions (PPI) are central to most cellular processes and, as such, are the focus of many probe- and drug-discovery programs. However, it has been difficult to identify small molecule or peptide inhibitors of PPI that bind stoichiometrically to a single site on the protein surface. Often sited reasons for this challenge include a) the flat nature of PPI interfaces, which sometimes lack deep grooves where small molecules can stick, b) the large contact area at the interface, which often exceeds the surface area of a drug-sized molecule, and c) bias in the screening libraries, which are selected by adherence to criteria – like the Rule of 5 (1, 2) – that might not suit PPI inhibitors. Furthermore, early screening approaches to PPI were prone to artifacts and tended to select hydrophobic compounds with non-drug-like mechanisms of action, such as aggregation-based inhibition or protein denaturation (3, 4, 5). Despite these challenges, there are now a number of drug-like inhibitors of PPI in the literature (5, 6, 7, 8). From these examples, we are beginning to develop “best practices” for selecting tractable targets, applying appropriate screening assays, and evaluating mechanisms of action. This chapter focuses on the selection and development of screening assays for identifying small molecules that can modulate PPI, and will touch on secondary assays used to remove artifacts and demonstrate binding. General introductions to assay development for HTS and to common assay equipment and instrumentation can be found in Basics of Assay Equipment and Instrumentation for High Throughput Screening.

The screening assays described here monitor the binding of the two proteins, and can be used to measure inhibition or augmentation of the PPI by small molecules or other modulators (e.g. antibodies, peptides). Table 1 provides a brief summary of the assays. All assays can be used for primary high-throughput screening (HTS) at single concentrations of test compounds and for dose-response assays to obtain IC50 values. These formats are regularly used in 96- and 384-well plates and can often be adapted to 1536-well plates. Traditional high-throughput assays involve binding one of the protein partners to the plate; here, we provide an overview of these ELISA-like assays, including DELFIA. Increasingly, screening scientists utilize mix-and-read assay formats such as fluorescence polarization/anisotropy (FP), fluorescent/Foerster resonance energy transfer (FRET, TR-FRET, HTRF), and bead-based assays like AlphaScreen (Perkin Elmer). These solution-phase assays are often easier to develop than ELISAs because they have fewer components, and are simpler and faster to run because they avoid multiple incubation/wash cycles. On the other hand, ELISA and DELFIA can be very sensitive and are still used for diagnostics and some high throughput screens.



Table 1: Overview of Assay Formats

General Considerations

Screening in general, and for PPI inhibitors in particular, identifies compounds with many mechanisms of inhibition. Some of these mechanisms are undesired, such as direct assay interference, nonspecific binding, and covalent modification. To reduce the likelihood of finding such compounds, we recommend the following practices for in vitro assays:


Use of detergents in biochemical assays. Low concentrations of detergents tend to stabilize proteins, reduce nonspecific binding of proteins to assay plates, and break up compound aggregates. We favor using one detergent in a screen, then following up with alternate detergents, since no one detergent removes all aggregates. Commonly used detergents include Triton X-100 (0.01%), Tween 20 (0.005%), and Chaps (0.1%); in general, detergents should be used at concentrations below their critical micelle concentration (CMC).


Use of carrier proteins. As with detergents, non-interfering proteins such as gamma globulin, casein, or Prionex (Centerchem) can be used to reduce nonspecific binding of compounds and assay proteins. Bovine serum albumin (BSA), used in most cell-based assays, binds many compounds that are viable drug leads and may not be the best choice for primary biochemical screens. Using carrier proteins is less common than detergents, but should be considered for primary or secondary testing.


Diluting compounds in DMSO. Most HTS libraries are dissolved in DMSO, and small concentrations of DMSO (typically <0.1%) are thus carried into assays. The sensitivity of an assay to DMSO must be determined during assay development. In addition, since compounds vary in their solubility, it is best practice to limit intermediate dilutions into buffer. In particular, when running dose-response titrations, the compounds should be serially diluted in DMSO, then transferred into the assay buffer as close to the final concentration as possible. The use of nanoliter volume dispensers has made this approach practical for 384-well and higher format plates.


Use of orthogonal assays. Compounds will interfere with assays in multiple ways, some of which are difficult to predict. It is therefore important to follow up any primary assay with an orthogonal secondary assay that uses a different modality (e.g. colorimetric vs luminometric) or format (plate-bound, vs mix-and-read, vs label-free). Finally, unless the goal is to discover covalent modifiers of a PPI, reversibility of inhibition should be demonstrated. This can be accomplished by mixing the reagents at high concentration, then diluting to a condition well below the affinity of the inhibitor. Additionally, the PPI partners can be incubated with inhibitor, then analyzed by mass spectroscopy.

ELISA-type assays


Enzyme-Linked Immunosorbant Assays (ELISAs) follow the basic design shown in Figure 1. These formats can be used to measure PPI and thus to measure competition between a PPI and a small-molecule inhibitor. To measure a PPI, one of the proteins is attached to a plate surface and the second protein is then allowed to bind to the first protein. The second protein is detected by binding of an antibody that is linked to an enzyme. When substrate is added, the enzyme produces a measurable readout that is quantitatively linked to the amount of the second protein. ELISAs can be very sensitive, because the readout is amplified by using an enzyme. Further amplification can be achieved by using multiple layers, such as secondary antibodies.

Figure 1: Format for ELISA and DELFIA assays


Figure 1: Format for ELISA and DELFIA assays. Left: An ELISA is built up in several steps, starting with antibody to protein 1, protein 1 (green), protein 2 (orange), anti-protein 2, and an anti-species antibody conjugated with an enzyme (AP = alkaline (more...)

ELISA technically means that the detection event uses an antibody and enzyme-based detection; however, this term is commonly used to describe plate-bound detection of a reagent, even if the affinity reagent is not an antibody or if the detection reagent is not an enzyme. One commonly used, non-enzymatic format is called Dissociation-Enhanced Lanthanide Fluorescent Immunoassay (DELFIA, Perkin Elmer). In DELFIA, the detection signal is time-resolved fluorescence of a lanthanide ion (such as europium). The lanthanide ion is bound to the affinity reagent through a chemical linkage; upon adding a proprietary detergent mixture, the europium fluoresces, providing a highly sensitive measurement of the concentration of bound protein. Three features of lanthanide fluorescence lead to highly sensitive and selective assays: a) a long emission lifetime (milliseconds) allows the measurement to start after the fluorescence of organic material (proteins, test compounds) has decayed, b) the emission occurs at around 600 nm, where few biological materials absorb or emit light, and c) the narrow emission spectrum of lanthanides allow them to be multiplexed.

ELISA-style assays can be designed in many ways. Attachment of the surface-bound protein can occur by passive adsorption to a plastic plate, by capture with an adsorbed antibody, or by biotinylation and avidin capture. Detection of the second protein can occur by direct labeling the protein with a signal-generating enzyme, by binding of an enzyme-labeled antibody, by binding of an unlabeled primary antibody followed by a labeled secondary antibody, or by biotinylation followed by enzyme-linked avidin. Detection enzymes can include colorimetric, fluorogenic, or luminogenic reactions. Selection criteria for of each of these steps are described below.

General Considerations

Consider the affinity of the PPI when designing a plate-based assay. Such assays involve multiple wash steps, which will remove unbound protein. If binding kinetics are rapid, as with most weak interactions (ca. > 1 μM), signals will be lower. The fewer amplification steps (e.g. direct conjugation of the enzyme to the solution-phase protein), the less signal is lost to washing. Assay format is therefore a compromise between assay complexity and signal amplification.

In principle, either member of the PPI could be immobilized, but several issues should be considered:


Is there a potential for avidity in the interaction? Avidity occurs when multiple contacts are made simultaneously, as with a trimeric protein binding to a trimeric ligand, or when a trimeric protein binds to a plate with a high density of ligand. This Velcro-like effect results in slowed unbinding kinetics (off-rates) and thus an apparent affinity that is tighter than the 1:1 binding affinity. For weak interactions, such avidity allows the PPI to survive washing, but it also complicates quantitative analysis. If one member of the PPI is monomeric, it is generally recommended to use this protein as the solution-phase protein, while a multimeric partner is immobilized on the plate.


Is one protein more likely to bind to the small molecule? Compounds are more likely to bind to the side of the PPI that is concave, such as when a bit of secondary structure from the other protein binds into a groove (examples include MDM2/p53, PDZ domains). If the grooved protein is kept in solution, then the ELISA will monitor a solution-phase binding event.


Is one protein more apt to precipitate or aggregate? If one protein is known to precipitate, it might be more stable as the captured, plate-bound partner.


Is one PPI partner limiting? Generally, more of the immobilized protein is used, so if one protein is easier to obtain and purify, it should be considered for immobilization.


Another option is if one of the proteins is expressed with a tag such as his or GST or FLAG, a plate coated with antibodies to the tag would serve to immobilize the protein to the plate. A protein can be biotinylated and streptavidin-coated plate can be used.

If there is no compelling reason to use one protein for surface-immobilization, then both orientations of the assay should be evaluated. In the ideal case, the same IC50 values should be obtained from both formats.

Assay design and development


Instrumentation: The assays described here can be performed with most multimodal plate readers, and the readout can be selected based on available instrumentation. Typical readouts are absorbance, fluorescence, luminescence, or time-resolved fluorescence (TRF). Time-resolved fluorescence, used by DELFIA, is the least-standard modality, but most HTS facilities will have TRF-compatible instruments.


Plates: ELISA-type assays are generally performed in polystyrene microwell plates with high-binding surfaces to adsorb proteins. The color of the plate is selected based on the readout: clear (absorbance), black (fluorescence), and white (luminescence). DELFIA can be performed in clear, white or yellow plates.


Binding of first protein (protein 1): The first immobilized protein can be bound to the plate by passive adsorption or by capture. Passive adsorption is simple – the protein is added to a bare plate; however, some proteins will denature upon adsorption and the orientation of proteins on the surface will be random. After the protein is adsorbed, the rest of the well surface is blocked with a nonspecific protein, such as casein (1%), nonfat milk (5%), or bovine serum albumin (1%). Critical steps for passive adsorption include a) selecting a concentration of protein to maximize ELISA signal and b) selecting a blocking protein that reduces nonspecific binding of the second protein partner and detection reagents. At any step in the ELISA process, a blocking protein or detergent can be used to reduce nonspecific binding.

If assay sensitivity is low when the protein is adsorbed, a capture step can be added. In this format, an antibody or avidin is adsorbed to the plate first. Generally, the capture protein is plated at a saturating condition. The plate is then blocked with nonspecific proteins and protein 1 is added. If an antibody is used for capture, it should not block the PPI and should be available in sufficient supply for the scale of the assay. If avidin is used, there are different versions (e.g. streptavidin and neutravidin) that could alter the degree of nonspecific binding. The protein to be captured must be biotinylated, which can be accomplished during expression (via AviTag; Avidity) or chemically (via reaction with amines, acids, or cysteine residues). Biotinylation using AviTag sequences will provide homogeneous labeling near the N- or C-terminus of the protein.


Binding of the second protein (protein 2): Protein 2 in the PPI can also be added to the ELISA plate in several formats. The protein can be unmodified, it can be biotinylated (if the first protein was not immobilized by biotin/avidin binding), or it can be labeled directly with the detector (e.g. enzyme or DELFIA probe). As mentioned above, the decision to label or use secondary detection is a balance between the number of washing steps and signal amplification. Directly labeled proteins, such as horseradish peroxidase (HRP)-protein conjugates, can be less soluble and more prone to artifacts; compounds can also interfere with detection, such as by inhibiting the detection enzyme itself. Therefore, ELISAs usually use a secondary detection step. In either case, protein 2 should be titrated to achieve a robust but non-saturating signal. Incubation times should allow equilibrium to be reached. After incubation, the unbound protein 2 should be thoroughly washed from the plate, usually using 3 cycles of phosphate-buffered saline with 0.01% Tween 20 (or other detergent).


Affinity reagents (e.g. antibodies or streptavidin): When protein 2 is not directly labeled with a detection reagent, one or two binding steps are required to add the detector. When protein 2 is biotinylated, avidin is conjugated with the detection reagent. When the protein is unmodified, an antibody to protein 2 is usually used. This primary antibody can be labeled with detection reagent or a secondary antibody (e.g. rabbit anti-murine IgG) can be labeled and bound. The affinity reagents should be titrated to reach a maximal signal-to-background; the binding can be saturated, but care should be taken to ensure that the reagent is not binding non-specifically to wells without protein 2. Incubation time is another optimize-able parameter, and can vary from 1 hour to overnight. After incubation, the plate should be thoroughly washed to remove unbound reagents.


Detection methodology: The final step of the ELISA involves adding a substrate; when the linked enzyme turns over the substrate, a measurable change (e.g. color) occurs. Two commonly used enzymes are horseradish peroxidase (HRP) and alkaline phosphatase (AP); avidin and antibody conjugates of these enzymes are widely available, as are chromogenic and luminogenic substrates. Generally speaking, luminescent substrates are more sensitive, requiring smaller amounts of material and/or less incubation time than chromogenic substrates and have a wider dynamic range than chromogenic substrates. On the other hand, chromogenic products tend to be more stable over time. ELISAs can be read kinetically (monitoring the color change over time) or at a fixed endpoint. Endpoint readings can include a quenching step, such as the addition of acid (e.g. equal volume of 1 M HCl or H3PO4) or base (e.g. 1 M ammonium chloride) to stop enzyme turnover and stabilize the ELISA signal.

DELFIA assays are processed similarly to ELISAs. Some europium-labeled reagents are commercially available, including anti-IgGs, anti-tag antibodies (such as anti-Histag antibodies), and streptavidin, and others can be prepared in the lab or by custom synthesis (for example, see Perkin Elmer and Cisbio). Detection requires the addition of a commercial “dissociation-enhancement solution.”

Benefits and limitations

ELISA and DELFIA have some important benefits. They are very flexible and sensitive, and can be inexpensive to run. They are also less likely to have compound interference since the compound is not present in the processing step. They also have significant limitations that have led to their reduced use in HTS. Most ELISAs have multiple incubation and washing steps, which are time-consuming for automated and bench-top assays. Washing can also disrupt weak interactions (e.g. Kd > 1 μM). Finally, it is important to demonstrate that potential PPI inhibitors are not interfering with the detection system or acting nonspecifically with the proteins and detection reagents. Changing formats and using complementary detection methods (such as those described below) will help to validate potential inhibitors.

Related technologies

PPI assays have also been performed with bead-based separation, e.g. using flow cytometry (9).

Mix-and-read assays

Three main formats are available with both similarities and differences with regard to label, secondary detection, and maximum distance allowable between the protein partners. FRET and AlphaScreen are proximity measurements, meaning that they rely on the protein partners being within 10 – 100 angstroms. Fluorescent Polarization measures the tumbling time, related to the molecular mass, experienced by a fluorophore. A principal advantage of these mix-and-read assay formats is that they requiring no washing steps, leading to a wider dynamic range and higher plate throughput.

Fluorescence polarization/anisotropy


Fluorescence polarization (FP) is a sensitive nonradioactive method for the study of molecular interactions in solution (10). This method can be used to measure association and dissociation between two molecules if one of the molecules is relatively small and fluorescent. When a fluorescently labeled molecule is excited by polarized light, it emits light with a degree of polarization that is inversely proportional to the rate of molecular rotation. Molecular rotation is largely dependent on molecular mass, with larger masses showing slower rotation. Thus, when small, fluorescent biomolecule, such as a small peptide or ligand (typically < 1500 Da), is free in solution, it will emit depolarized light. When this fluorescent ligand is bound to a bigger (e.g. > 10, 000 Da) molecule, such as a protein, the rotational movement of the fluorophore becomes slower and thus the emitted light will remain polarized. Thus, the binding of a fluorescently labeled small molecule or peptide to a protein can be monitored by the change in polarization (Figure 2).

Figure 2: Diagram of a fluorescence polarization assay


Figure 2: Diagram of a fluorescence polarization assay. Rapidly rotating small molecule fluorophore gives low FP signal (low mP). The association of a relatively large molecule, such as a protein, with the small molecule fluorophore slows down the motion (more...)

Assay design


Selection of FP probe: Protein-protein interactions can be monitored by FP if one of the components of the PPI is small. Typically, the molecular weight of the ligand/probe is less than 1500 Da, although up to 5000 Da can be acceptable if the binding partner is very large. For most PPI, FP will be practical only a) if one side of the PPI can be minimized to a peptide, b) if there is a synthetic peptide known to bind at the interface (e.g. via phage display), or c) if an organic compound binds at the interface (or to a mutually exclusive binding site). Fortunately, there are several examples of peptides that mimic the epitope of a protein in a PPI, including PDZ domains, IAPs, Bcl2-family proteins, and others.


Selection of fluorescent dye: Once a probe molecule has been selected, it must be labeled with a fluorescent dye. Dyes are typically available in amine-reactive, cysteine-reactive, and acid-reactive forms and are chemically attached to the probe peptide/molecule using simple chemistry. Typical fluorophores used in FP are fluorescein, rhodamine, and BODIPY dyes. The BODIPY dyes have longer excited- state lifetimes than fluorescein and rhodamine, making their fluorescence polarization sensitive to binding interactions over a larger molecular weight range (11). Red-shifted dyes are preferable to reduce the number of compounds that will cause interference with the 405nm (e.g. fluorescein) range.


Selection of buffer: the buffer must have low fluorescence background. Frequently used buffers have neutral pH such as PBS, HEPES.


Instrumentation for FP measurement with microtiter plate: Many commercially available instruments are capable of measuring the FP signal from solution in 96/384/1536-well microtiter plate format for high throughput screening (HTS). The fluorescence is measured using polarized excitation and emission filters. Two measurements are performed on every well and fluorescence polarization is defined and calculated as:

Polarization = P = (Ivertical – Ihorizontal)/(Ivertical + Ihorizontal)

Where Ivertical is the intensity of the emission light parallel to the excitation light plane and Ihorizontal is the intensity of the emission light perpendicular to the excitation light plane (10). All polarization values are expressed as the milli-polarization units (mP).

All commercial microplate readers have built-in software for mP calculation. Depending on the instrument used, three sets of data are generally reported, including calculated mP values, raw fluorescence intensity counts of vertical (or Parallel/S-channel) and horizontal (or perpendicular/P-channel) measurements for each well.

mP calculation for different instruments requires the proper use of measured fluorescence intensity of parallel/S-channel and perpendicular/P-channel. As optical parts of fluorometers possess unequal transmission or varying sensitivities for vertically or horizontally polarization light, such instrument artifacts should be corrected for accurate calculation of the absolute polarization state of the molecule using fluorescent readers. This correction factor is known as the "G Factor” which is instrument-dependent. G-factor corrects for any bias toward the horizontal (or perpendicular/P-channel) measurement. Most commercially available instruments have an option for correcting the single-point polarization measurement with G factor. For example, the mP values for FP measurement with Envision Multilabel plate reader are calculated as:

mP = 1000 * (S - G * P) / (S + G * P)

In practice for HTS applications, however, it is unnecessary to measure absolute polarization states; the assay window is what is important. The assay window is insignificantly changed by G Factor variation.


Determining the concentrations of fluorescent probes for the FP binding assay: In order to select the proper concentrations of fluorescent probe for the binding assay, increasing concentrations of fluorescent probe is prepared in assay buffer without the binding protein. The fluorescence intensity (FI) in the parallel channel is then measured with defined settings in a plate reader with FP mode. A concentration of the fluorescent probe with at least 10-fold or higher FI signal compared to that of buffer only should be selected for the subsequent binding assay. Notice that the FP signal is expressed as a ratio of fluorescence intensities. Thus, the signal is not influenced by changes in intensity brought about by changes in the tracer concentration.


FP Binding assay development: To determine the binding of the fluorescent probe with the protein of interest, increasing concentrations of protein are mixed with a fixed concentration of the probe. The FP signal as expressed in mP is then measured with a plate reader. mP vs [protein] is then plotted to generate a binding isotherm for the calculation of association parameters such as Kd and maximal binding. For the FP inhibition assay, select a concentration of protein that provides ca. 80% of the maximum change in polarization for the probe (e.g. 80% bound).


Data analysis: The dynamic range of the FP assay, i.e. assay window, is defined as mPb – mPf, where mPb is recorded mP value for the specific binding in the presence of a particular protein concentration and mPf is the recorded mP value for free tracer from specific binding proteins (12). Typically, the assay window is 3-5 fold (e.g. 50 mP – 150 mP).


Selectivity: once the concentrations of probe and protein are determined, the specificity of the interaction should be assessed. First, an unlabeled version of the probe is titrated into a mixture of the FP-probe and protein. As the concentration of unlabeled probe competes with bound fluor-probe, the FP should decrease. The IC50 for this interaction should be similar to the Kd measured above. Similarly, other known inhibitors should yield the expected IC50 values.


Fluorescence of the bound probe: often the fluorescence of the probe changes when it binds to the protein. In this case, anisotropy measurements should be used in place of polarization, since unlike FP, anisotropy is directly proportional to fluorescence intensity. Anisotropy is calculated with the following expression:

Anisotropy: r = (Ivertical – Ihorizontal)/( Ivertical + 2 Ihorizontal)

where Ivertical is the intensity of the emission light parallel to the excitation light plane and Ihorizontal is the intensity of the emission light perpendicular to the excitation light plane.

Case study: Monitoring 14-3-3 protein interactions with a homogeneous FP assay

The 14-3-3 proteins mediate phosphorylation-dependent protein-protein interactions. Through binding to numerous client proteins, 14-3-3 controls a wide range of physiological processes and has been implicated in a variety of diseases, including cancer and neurodegenerative disorders (13). We have designed a highly sensitive fluorescence polarization (FP)-based 14-3-3 assay (Figure 3), using the interaction of 14-3-3 with a fluorescently labeled phosphopeptide from Raf-1. The specificity of the assay has been validated with known 14-3-3 protein antagonists, e.g., R18 peptide, in a competitive FP assay format. The signal-to-background ratio is greater than 10 and a Z’ factor is greater than 0.7 (12). Because of its simplicity and high sensitivity, this assay is generally applicable to studying 14-3-3/client protein interactions and for HTS.

Figure 3: Development of FP assay for TMF-pS259-Raf/14-3-3


Figure 3: Development of FP assay for TMF-pS259-Raf/14-3-3. A: The interaction of 14-3-3 with TMR-pS259-Raf gave rise to a significant FP signal with a minimal background polarization with the peptide probe alone or with increasing concentrations of a (more...)


Protein (14-3-3γ): the recombinant GST-14-3-3 protein was expressed in Escherichia coli strain BL21 (DE3) as a GST-tagged product and purified


Probe (TMR-pS259-Raf): a phosphopeptide derived from Raf-1 was synthesized and labeled with 5/6 carboxytetramethylrhodamine (TMR)


Buffer: HEPES buffer containing 10 mM HEPES, 150 mM NaCl, 0.05% Tween 20 DTT 0.5 mM DTT, pH 7.4


384-well black plate (Corning Costar Cat#: 3573)


Instrument for FP measurements: FP measurements were performed on Analyst HT plate reader (Molecular Devices, Sunnyvale, CA) using FP protocol. For Tetramethylrhodamine (TMR)-labeled probe (Excitation: 545 nm; Emission: 610 nM), a dichroic mirror of 565 nm was used.


Selection of probe concentration: 1 nM of the TMR-pS259-Raf peptide was chosen for the binding assay based on the observation that 1 nM of the TMR-labeled peptide exhibited about 10 times more fluorescence intensity in the parallel channel than the “buffer-only” control samples.


Prepare probe and protein solutions: 2 basic solutions were prepared. Solution A contained TMR-pS259-Raf peptide in the HEPES buffer (2× solution with 2 nM of the peptide probe or as specified). Solution B contained 14-3-3 proteins in HEPES buffer (2× solution with increasing concentrations of 14-3-3 protein; a serial dilution approach is generally used for the protein titration).


Binding FP assay: The 14-3-3 FP binding assay was carried out in black 384-well microplates in a total volume of 50 µl in each well. For each assay, a 25 µl of Solution A (probe) is mixed with 25 µl of solution B (protein) in 384-well plate. Probe only without protein is always included as blank control.


FP measurement: the polarization value in mP was measured at room temperature (RT) with an AnalystHT reader immediately, or after incubating at RT as desired time period for the equilibration of the interaction.


Competitive FP assays: the specificity of the FP binding assay is generally evaluated with known antagonists, e.g, unlabeled probe, peptide or small molecule antagonists, in competitive FP assay. To achieve the desired sensitivity, the concentrations of fluorescent Raf peptide probe and 14-3-3 protein are carefully chosen to maximize the difference between the highest and lowest polarization values. Serial dilutions of competitive peptide (R18) are added to a reaction buffer containing TMR-pS259-Raf (1 nM) and GST-14-3-3γ (0.5 µM) and incubated at RT for 1 hr. The mP values were measured and the competitive effect was expressed as percentage of control mP (TMR-pS259-Raf and GST-14-3-3γ) after subtracting the background mP (TMR-pS259-Raf alone).

Benefits and limitations:

FP-based technology has a number of key advantages for monitoring bimolecular interactions, especially for HTS applications. It is nonradioactive and is in homogenous “mix-and-read” format without wash steps, multiple incubations, or separations. FP measurement is directly carried out in solution; no perturbation of the sample is required, making the measurement faster and perhaps more native-like than immobilization-based methods like ELISA. It is readily adaptable to low volume (30 µl for a 384-well plate or 5 µl for a 1536-well plate). In addition to measuring PPI, FP assays have been used to study a wide variety of targets including protein-nucleic acid interactions, kinases, phosphatases, proteases, G-protein-coupled receptors (GPCRs), and nuclear receptors (14, 15).

As a fluorescence-based technology, FP is subject to optical interference from compounds that absorb at the excitation or emission wavelengths of the fluorescent probe. Being a ratiometric technique makes FP somewhat resistant, though enough light must be available to obtain an emission signal. FP is also sensitive to the presence of fluorescence from test compounds. The use of red-shifted probes will minimize background fluorescence interference.

Fluorescent/Förster resonance energy transfer and time-resolved (TR) FRET


Fluorescence/Förster Resonance Energy Transfer (FRET) is the phenomenon of non-radiative energy transfer between two fluorophores with specific spectral properties. In order for FRET to occur, the emission spectrum of one fluorophore, i.e. the “donor,” must overlap the excitation spectrum of the second fluorophore, i.e. the “acceptor.” When the donor is excited by incident light, energy can be transferred to the acceptor via long-range dipole-dipole interactions, resulting in acceptor emission; however, this FRET event will only occur if the donor and acceptor are in sufficient proximity to one another. FRET efficiency E is defined by the equation

Image ppi_eq1.jpg

where r is the distance between the fluorophores and Ro is the Förster distance and which FRET efficiency is 50% for the specific donor/acceptor pair. Two key factors arise from this equation. First, the amount of energy transfer decays with the sixth power of the distance between the fluorophores. Second, the term Ro depends on the spectral overlap of the donor emission spectrum and the acceptor absorbance spectrum; FRET can be observed over longer distances when the spectral overlap is large. Fortunately, the proximity limit for several donor/acceptor pairs is approximately 10 nm, which happens to be the distance over which many biomolecular interactions occur. Therefore, FRET can be used to monitor biomolecular interactions in a homogeneous mix-and-read assay format by tagging or labeling interacting biomolecules “A” and “B” with acceptor and donor fluorophores, respectively. In such a scenario, the ratio of acceptor to donor emission following donor excitation is used to quantify and monitor “AB” binding. Table 2 lists some common donor/acceptor pairs.



Table 2: Common donor/acceptor pairs for FRET and TR-FRET/HTRF1

Due to the spectral properties of biological media and traditional FRET donor/acceptor pairs, the FRET signal can be significantly contaminated by 1) autofluorescence of biological media and test compounds; 2) a wide acceptor excitation spectrum that allows the acceptor to be directly excited by incident light; and 3) a wide donor emission spectrum that bleeds through into the acceptor emission detection window. These signal contaminants must be corrected for and can significantly diminish the sensitivity of traditional FRET assays.

One elegant solution to the problem of FRET signal contamination is the use of donor fluorophores with exceptionally long emission half-lives (up to 1500 µs), such as the rare earth metals Europium or Terbium, in a modification of FRET known as Time Resolved (TR) FRET (also called HTRF). In TR-FRET, Europium or Terbium cryptates (ligands that coordinate the metal ion and provide an “antenna” dye) serve as donors that have a very long luminescence half-life. This long emission decay allows for a time delay (50-150 µs) between donor excitation and the recording of acceptor emission. During this time delay, both media autofluorescence and acceptor excitation due to incident light will rapidly decay (ns scale) and be extinguished by the time acceptor emission is measured. This essentially eliminates signal contaminants 1 and 2 above. Signal contaminant 3 – donor emission bleedthrough into acceptor detection – is attenuated by the use of acceptors with red-shifted emission (Table 2) such as allophycocyanin, Alexa 680 (Invitrogen), Cy5, or d2 (Cisbio Bioassays). Another advantage of TR-FRET is that the rare earth metals have a modestly larger proximity limit for FRET (up to 20 nm), allowing for the detection of larger biomolecular complexes. TR-FRET assays are well suited for certain HTS applications due to their homogenous mix-and-read design, high signal-to-background ratios, and enhanced proximity detection range (Figure 4).

Figure 4: Principles of TR-FRET


Figure 4: Principles of TR-FRET. A: Schematic of a typical FRET bioassay. Protein 1 is bound to an antibody fused to a donor fluorophore, e.g. Terbium (Tb), and Protein 2 is bound to an antibody fused to an acceptor fluorophore, e.g. d2 or XL665. If A (more...)

Assay Design

Instrumentation: A plate reader capable of allowing a time delay between excitation and fluorescence detection is required. The multimodal readers Envision (PerkinElmer) and Analyst HT (Molecular Devices) and PheraStar (BMG) are well suited for this application.

2. Plates: TR-FRET assays are performed in black opaque plates. Assays may be performed in 96- 384-, 1536-well formats.

3. Buffers: The following buffer is used routinely at the Emory Chemical Biology Discovery Center for all TR-FRET assays: 20 mM Tris, pH 7.5, 0.01% Nonidet P40, and 50 mM NaCl. However, multiple buffers can be used. As noted in the general considerations above, the use of detergents (e.g. Nonidet P40, Triton X-100, Tween 20) and carrier proteins (e.g. - Prionex) should be optimized during assay development to reduce nonspecific effects. Similarly, salt conditions can affect PPI.

4. Labeling Reagents: Proteins can be directly labeled with FRET donor and acceptor, using amine-, acid-, or cysteine-reactive dyes. Because random chemical coupling can disrupt the PPI, it is more common to use anti-epitope tag antibodies or streptavidin labeled with FRET and TR-FRET dyes. These general protein-dye conjugates are commercially available for proteins containing GST, HA, Flag, 6xHis, myc, or biotin tags. If it is necessary to measure binding of untagged or endogenous biomolecules, kits for labeling primary (anti-protein) antibodies with FRET and TR-FRET fluorophores are also available for purchase. FRET and TR-FRET reagents are available from Cisbio Bioassays, PerkinElmer, and Invitrogen, among others.

5. Assay Conditions: FRET and TR-FRET assays are performed at room temperature. Assay performance has been shown to be stable for up to 24 hours at room temperature. In some formats, the quality of the signal improves with time and can be much better after overnight incubation. The timing for signal development and decay should be confirmed for each assay.

Steps for developing a TR-FRET Assay

1. Select binding partners to be used. Typically, the greatest TR-FRET sensitivity is obtained when the purified, recombinant protein-binding domains of interacting proteins are used. However, if the binding domains are not known, full-length recombinant proteins can also be used. Additionally, if purified proteins cannot be generated, or if it is critical to evaluate the PPI in a complex milieu, TR-FRET can also be performed using cell lysates containing over expressed, epitope-tagged versions of the interacting proteins. If using cell lysates, it may be best to develop stable cell lines expressing one of each binding pair to ensure consistent protein expression.

2. Select protein concentrations. When proteins are directly labeled with fluorophores, simply titrate each binding partner in a matrix to determine concentrations that yield optimal assay window, signal-to-background ratio, and Z’ values. When proteins are not directly labeled, the concentrations of the PPI and the dye-conjugated antibodies/avidin must also be optimized. It is typical to start with constant concentrations of the dye-conjugated antibody/avidin, and titrate the PPI partners. Most commercial reagents suggest starting conditions; in general, the concentrations of FRET-conjugate antibodies should be higher than the concentrations of the proteins they detect (e.g. 20 nM anti-HA antibody and 10 nM HA-tagged protein).

3. Select concentrations of (TR-)FRET reagents. Once the PPI concentrations have been selected, the FRET-tagged antibodies/avidin should be titrated to optimize the assay window, signal-to-background, and Z’ values. When concentrations of antibodies are too high, the efficiency of FRET can decrease. This effect, called “hooking,” is described in the General Consideration: Hooking Effect section below.

4.Test effect of DMSO. Assay performance should be measured over a range of DMSO concentrations to ensure that screening results are not skewed by vehicle effects. For HTS, DMSO generally ranges from 0.1 – 1%, but higher levels are sometimes acceptable.

5. Assess assay performance. Positive controls are then titrated in a competition format to ensure that IC50 values match expectation. Either known interaction inhibitors or non-labeled binding partners can be used as positive controls for binding competition/disruption.

Example: Performing a TR-FRET Assay


All assay components are combined with assay buffer (e.g. 20 mM Tris, pH 7.5, 0.01% Nonidet P40, and 50 mM NaCl) to their optimized concentrations and 19 µL are transferred to each well of a 384-well plate.


Test compounds are added. In this case, test compound stocks are at 1 mM, and 0.5 µL is added to each well to give a final compound concentration of 25 µM.


All assay plates must contain at least one column of minimal FRET/background control (e.g., all assay components minus one binding partner, usually the one that binds the acceptor fluorophore) and at least one column of maximal FRET vehicle control (i.e., all assay components plus DMSO).


The plate is incubated at room temperature for one hour or overnight and then the FRET signal is recorded.


Background is subtracted from all FRET values and test compounds are compared to the maximal FRET control to determine percent inhibition of binding for each compound.

Benefits and Limitations

Several factors make FRET and TR-FRET attractive techniques to measure PPIs. As with the other mix-and-read formats, FRET methods are relatively easy to automate and to miniaturize. The approach is also flexible, since many dyes and dye-antibody conjugates are available. In contrast to FP, FRET can be used with a wide range of protein sizes, with the proviso that the FRET pairs must come within a few nanometers of each other. Thus, TR-FRET assays can be performed with peptides, full-length recombinant proteins, transfected cell lysates, and, in some cases, with endogenous proteins in cell lysates. This potentially allows for the development of robust HTS screening assays using binding pairs in a less artificial environment. FRET and TR-FRET are usually performed as ratiometric assays, which reduce the effects of autofluorescence and spectral interference of media and test compounds. TR-FRET further reduces the effect of autofluorescence by allowing organic fluorescence to decay before TR-FRET is measured. Finally, TR-FRET formats allow multiplexing. For instance, FRET acceptors can be multiplexes with a single lanthanide donor, allowing two or three pairs of PPI to be monitored in a single well (16, 17). TR-FRET has been multiplexed with FP, providing increased information in the primary HTS screen (18).

There are limitations to the FRET and TR-FRET formats, however. The signal window for FRET experiments depends on several factors implicit in the Forster equation, including the orientation of the dyes and the size of the complex – including the size of the FRET-labeled antibodies. For very large complexes, AlphaScreen (see AlphaScreen Format) could yield a stronger signal. Furthermore, while TR-FRET’s ratiometric format does reduce interference from test compounds, those compounds that absorb a lot of UV light can inhibit excitation of the FRET donor, which absorbs in the far-UV (ca. 350 nm). Finally, if a test compound interferes with the binding of the fluorophore-tagged antibodies to their epitopes it will be detected as a hit in a TR-FRET screen, even though it has no effect on the binding of the target molecules themselves. Following up with controls and secondary assays will remove such compounds from consideration.

AlphaScreen Format


AlphaScreen™ is bead-based format commercialized by PerkinElmer ( and used to study biomolecular interactions in a microplate format. A newer, more sensitive version of the technology is called AlphaLISA. The acronym ALPHA stands for Amplified Luminescent Proximity Homogeneous Assay. The technology of AlphaScreen was originally developed under the name LOCI® (Luminescent Oxygen Channeling Immunoassay) by Dade Behring, Inc. of Germany (19). Like FRET, AlphaScreen is a non-radioactive, homogeneous proximity assay. Binding of two molecules captured on the beads leads to an energy transfer from one bead to the other, ultimately producing a fluorescent signal. Excitation of the donor bead leads to the formation of singlet oxygen, which diffuses to the acceptor and stimulates emission. Unlike FRET, acceptor emission occurs at a higher energy (lower wavelength) than donor excitation.

The AlphaScreen assay beads are latex-based and approximately 250 nm in diameter. Both bead types (Donor and Acceptor) are coated with a hydrogel that minimizes non-specific binding and self-aggregation and provides reactive aldehyde groups for conjugating biomolecules to the bead surface. The beads are small enough that they do not sediment in biological buffers and bead suspensions do not clog the tips used commonly in liquid handling devices. The beads are typically used at ug/mL concentration and are very stable, even if heated to 95°C for example, for PCR, or lyophilized.

Donor beads contain a photosensitizer, phthalocyanine, which converts ambient oxygen to an excited form of O2, singlet oxygen, upon illumination at 680 nm. Like other excited molecules, singlet oxygen has a limited lifetime prior to returning to ground state. Within its 4 μsec half-life, singlet oxygen can diffuse approximately 200 nm in solution. If an Acceptor bead is within that distance, energy is transferred from the singlet oxygen to thioxene derivatives within the acceptor bead, resulting in light production. Without the interaction between donor and acceptor bead, singlet oxygen falls to ground state and no signal is produced. AlphaScreen Acceptor beads use rubrene as the final fluorophore, emitting light between 520 and 620 nm. AlphaLISA acceptor beads use a Europium chelate as the final fluorophore, emitting light in a narrower peak at 615 nm (Figure 5). The AlphaLisa light is less likely to be affected by particles and other substances commonly found in biological samples (for example, plasma and serum), thereby reducing background noise and optimizing precision.

Figure 5: AlphaScreen and AlphaLisa


Figure 5: AlphaScreen and AlphaLisa. Left: Binding of biological partners (represented by small ovals A and B) brings Donor and Acceptor beads (represented by the large blue and yellow circles) into close proximity (≤200 nm) and thus a fluorescent (more...)

AlphaScreen assays have been developed to quantify enzymes, molecular (protein, peptide, small molecule) interactions, as well as DNA and RNA hybridizations. Due to the large diffusion distance of singlet oxygen, the binding interactions of even very large proteins and phage particles can be quantified by AlphaScreen and AlphaLISA. The high sensitivity and large distance range have led to increasing use of these technologies in HTS settings.

General Consideration: Hooking Effect

The hook effect is a common phenomenon found when using any sandwich-type assay, including AlphaScreen, ELISA, and some of the FRET-based formats described above. When the PPI components are titrated (e.g. during assay development), both donor and acceptor beads become progressively saturated by their target molecules, and the signal increases with increasing protein concentration. At the “hook” point, either the Donor or the Acceptor component is saturated with the target molecule and a maximum signal is detected. Above the hook point, there is an excess of target molecules for the donor or the acceptor beads, which inhibits their association and causes a progressive signal decrease (Figure 6). When the affinity of the PPI is higher (weaker) than the concentrations used in the assay, the hooking effect can be masked, resulting in what looks like a traditional saturation curve that reaches a plateau, rather than hooking. In this case, two competing equilibria are occurring: the signal is decreasing because of the hooking effect on the bead, but the protein-protein interaction is still being increasing because higher concentrations of protein drive the equilibrium toward more protein-protein complex. In either event, choose a protein concentration below the hook point (or saturation point) for your assay.

Figure 6: Hooking Effect in AlphaScreen


Figure 6: Hooking Effect in AlphaScreen. These principles hold for all sandwich-based assay formats (adapted from PerkinElmer).

Assay Design and Development


Instrumentation: Specialized instrumentation is required to read AlphaScreens since a high-energy laser is needed to excite the donor. However, most major companies who manufacture plate readers now provide models with AlphaScreen capability. Multimode readers suitable for AlphaScreen/AlphaLisa include PerkinElmer’s Envision and Enspire, Biotek’s Synergy, BMG’s PheraStar, FluoStar, and PolarStar, and Berthold Technologies Mithras LB 940. Specialized Alpha readers include PerkinElmer’s AlphaQuest and FusionAlpha.


Plates: AlphaScreen assays are performed in white opaque plates. Assays may be performed in 96-, 384- or 1536-well formats. Some plates, such as the ProxiPlate (PerkinElmer), have been optimized for AlphaScreen to place the sample closer to the detector and therefore give an increased signal. Excitation and signal measurement are both accomplished from the top of the plate. The measured signal is in part dependent upon reflected light, therefore the reflective properties of the plate influence signal. Higher density plates (e.g. 384- vs 96-well, 1536- vs 384-well) generally provide more sensitive AlphaScreen assays. First, the higher density wells are narrower and more efficiently reflect the emitted light back to the detector. Second, higher density plates allow a higher proportion of the sample to be excited by the 1 mm laser beam, leading to proportionately greater signal generation. Higher density plates also allow less reagent use, lowering the overall cost of the screen.


Buffers: Choose pH buffering capacity and salt concentration that will facilitate the desired interactions between the components of the assay. The following buffers have been used without problems: Acetate, HEPES, Bis-Tris, MES, Bis-TRIS propane, MOPS, CAPS, Phosphate, Carbonate, PIPES, Citrate, Formate, and Tris. pH values between pH 2.5 to 9 are well tolerated. Higher pH is also tolerated but there may be some loss of signal. If metal co-factors are needed for the PPI, it is best to titrate these components appropriately, but note that high concentrations will quench the signal; in particular, Al2+, Fe2+, Fe3+, Cu2+, Ni2+ and Zn2+ have been shown to quench singlet oxygen in the mM and sub-mM ranges (100 μM for Fe2+).

Detergents and/or blocking proteins should be used to reduce non-specific binding (see General Considerations). For most AlphaScreen applications, a BSA concentration of 0.1% (w/v) is sufficient to minimize non-specific interactions; alternate blocking reagents such casein, gelatin, heparin, poly-lysine, salmon sperm DNA, or Dextran T500 can be used (see Assay Design). The preservative azide can act as a potent scavenger of singlet oxygen and will inhibit the AlphaScreen signal, so Proclin 300 (Sigma-Aldrich) is recommended as a preservative and anti-microbial agent.


Kits: Generic detection kits from PerkinElmer include pre-coated beads that capture biotinylated, FITC-labeled, DIG-labeled, GST-tagged, 6X His-tagged, Protein A, Protein G, Protein L and anti-species beads. Unconjugated Donor and Acceptor beads are also available for direct conjugation of an antibody or other reagent of choice. Note that if you have purified your GST-tagged proteins or His-tagged proteins using an affinity column and will be using a GSH or Ni2+ bead in your Alpha assay, you will need to dialyze away any glutathione or imidazole in your purified protein preparation. These components will interfere with the interaction between the tagged protein and the bead. PerkinElmer also sells specific kits for various applications.


Titration of reagents: It is important to optimize the concentrations of each protein conjugated to the Donor and Acceptor beads. On the one hand, the amount of PPI formed is dependent on the concentrations of each protein and the affinity of the interaction. Until saturation is achieved, increasing the concentration of either protein will push the equilibrium towards higher complex formation. On the other hand, each type of Alpha bead has a specific binding capacity; once the beads are saturated with associated protein, additional protein may lead to a hooking effect (see above).

Binding capacities are influenced by a number of factors, including the size of the protein and the affinity of the bead for the protein. First, there is usually a higher binding capacity for smaller proteins. For instance, streptavidin-coated beads at 20 μg/mL usually saturate at around 30 nM of biotinylated peptide (ca 1.5 KDa), but saturate at around 2-3 nM of biotinylated antibody (ca 150 KDa). Second, the saturation point of a bead varies depending on its affinity reagent. For instance, anti-GST antibody beads bind more tightly to GST-labeled proteins than do glutathione beads. Hence, the saturation point is usually 20 nM of GST tagged protein binding to anti-GST beads, but 200 nM of GST-protein binding to glutathione-conjugated beads. More information on capacity can be found at the Perkin Elmer website (


Assay Conditions: The beads are sensitive to light and temperature. It is important to store the beads in the dark and conduct the parts of the assay that include the beads in low light conditions (less than 100 Lux) or to use green filters. Affected areas of the lab include the bench, plate reader and liquid handler. Finally, the chemistry is designed to give best results at room temperature (e.g.: 20–25°C); do not chill plates or incubate on ice before reading. Typically the AlphaScreen signal variation is 8% per °C so consistent temperature is important.

Steps for developing the assay:


Choose a suitable buffer system, noting the boundaries described above.


Titrate each binding partner to ascertain the optimal concentrations. For initial experiments, a final bead concentration of 20 μg/ml is recommended for both Donor and Acceptor beads. Subsequent dilution of the beads may be assessed once it is known that a sufficiently high signal/background can be achieved. Typically, most AlphaScreen assays will utilize a final concentration of biotinylated binding partner in the nanomolar range (e.g.: 0.5 nM–30 nM with 20 μg/mL of beads). Concentration ranges for each binding partner that interact directly with a capture molecule on the AlphaScreen beads vary considerably (ex.: 0.1 nM–300 nM) depending on the affinity of the binding partners, the efficiency of labeling, and/or stoichiometry of the capture tag/epitope.


It may also be necessary to vary the order of addition of the components to permit the most efficient interactions.


Incubation times need also to be optimized.

Benefits and limitations

Alpha technologies have become popular in recent years, likely because they are adaptable to many assay types, are very sensitive, and are active over long distances (200 nm vs 10-20 nm for TR-FRET). The central limitations to AlphScreen and AlphaLisa are the increased expense vs other mix-and-read formats and the sensitivity of the materials to ambient light.

Glickman, et al (20), compared FRET, TR-FRET and AlphaScreen formats (20), and concluded that the ALPHAScreen format had the best sensitivity and dynamic range. Of the three formats, TR-FRET assay had the least inter-well variation, most likely because it is a ratiometric type of measurement. Both FRET-type and AlphaScreen formats can measure a wide range of affinities (Kd‘s ranging from low pM to low mM) because there are no wash steps. It is noteworthy that AlphaScreen beads have 300-3000 proteins/bead, and the protein density can be varied. This multi-valency can significantly increase sensitivity, because one PPI per bead pair leads to the maximal signal. Furthermore, high concentrations can cause avidity. On the one hand, avidity augments the apparent binding affinity of the PPI, so less material is required. On the other hand, avidity might not be desired (e.g. for high affinity PPI or high affinity inhibitors), and the apparent IC50 values for inhibitors could be significantly weaker than the actual affinity of the inhibitor/protein interaction. With FRET methods, each binding partner carries a single label so that if some beads are unbound, a lower signal will be generated. However, monovalency also implies that the signal will be proportional to the number of binding interactions.

Validating drug-like binding of PPI inhibitors

The goal of primary screening is to select a set of compounds that might be active. Among “actives,” however, are many compounds that act by mechanisms that will not be optimizable into a qualified drug lead or biological probe. Some of the artifactual mechanisms that lead to activity in a primary assay are specific to the assay format; as described above, compounds could autofluoresce or quench the fluorescence signal used to detect the PPI. Thus, it is very valuable to develop at least one orthogonal assay formats, or an in vitro assay and a cell-based assay, plus an independent way to measure binding directly.

Other artifacts are less selective to the assay methodology, though they may be somewhat selective for the proteins or assay conditions. A well-described and very common example is compound aggregation (3). Aggregates can be quite large (30-200 nM) and can interfere with protein structure in a number of pathological ways. Aggregation can also be very dependent on the assay condition; rather than thinking of compounds as “aggregators,” it is more accurate to think of aggregation as a form of molecular interaction, dependent on salt, pH, detergents, carrier proteins, and concentration of the compound. Thus, it is not sufficient to demonstrate that a compound is selective for a particular screen over other screens; to be a bona fide PPI inhibitor, the compound must bind at a distinct site(s) on one of the proteins in the complex. Binding stoichiometry is therefore a key metric for selecting useful and optimizable probes/leads.

There are a number of biophysical assays that measure binding of the small molecule to the protein. It is very beneficial to use at least two assays, since no assay is infallible, and different types of information can be gleaned from each format. Depending on the size of the protein(s), the binding affinity of the molecule, and other details, the following methods can be used:

Optical Biosensors: There are several related technologies for measuring the binding of a surface-immobilized “ligand” to a soluble “analyte.” In general, optical biosensors detect changes in the angle, color, or phase of light reflected off of a solid/liquid interface. Many instruments are sensitive enough to monitor the binding of a small-molecule analyte to a surface-bound protein. These systems can also be used in competition experiments, in which a PPI is monitored in the presence of increasing concentrations of inhibitor. Because the signals are proportional to the change in mass of the analyte, PPI are usually easier to monitor than protein/small-molecule interactions.

The first popular optical biosensor was the surface plasmon resonance (SPR) instrument developed by Biacore (GE Healthcare). The technology is now widely used, and numerous companies market SPR instruments (e.g. Bio-Rad, ICX, and others). Most SPR instruments use microfluidics to introduce the analyte, and monitor the binding in real time. The concentration of analyte can be varied to develop a dose-response. Through kinetic and/or steady-state experiments, SPR provides a measure of binding stoichiometry, reversibility, and affinity to a protein bound to a surface. For recent descriptions of how to analyze and evaluate small-molecule SPR data, see Rich et al, 2011 and Gianetti et al, 2008 (21, 22).

Other technologies include optical gradients (SRU BIND, Corning Epic) and interferometry (Forte Bio Octet Red). The optical gradient systems are plate-based, allowing high throughput, but more limited kinetic resolution. In interferometry, the ligand is coated onto fiber optic sensors that are then dipped into solutions of analyte. This technology is developing rapidly, and could soon have the throughput, cost/assay, and sensitivity to rival SPR for measuring small-molecule/protein interactions.

Nuclear Magnetic Resonance (NMR): NMR measures the response of nuclei in a magnetic field, and is very sensitive to the chemical environment of the nucleus. Due to the flexibility of the method, NMR has many uses in small-molecule characterization and protein structure determination. Small-molecule/protein NMR experiments come in two general formats – ligand-detected and protein-detected. Ligand-detected experiments measure the change in the compound’s NMR signals (“resonances”) as a function of binding to protein. Energy can be transferred from the solvent and/or protein to the compounds (Saturation Transfer Difference, WaterLOGSY) or the apparent mass of the compound can be increased due to binding a large protein (translation, diffusion). Ligand-detected measurements are often used qualitatively, to assess the presence of binding to the target. Saturation Transfer Difference is particularly popular for moderate-throughput applications, because the protein concentration is low (micromolar) and it is particularly effective for compounds in fast exchange (weaker than micromolar). There is no limit on the protein size for ligand-detected experiments.

Protein-detected NMR provides a measurement of the effect of the compound on the protein. The most popular moderate-throughput methods are 15N-1H HSQC and 13C-1H HSQC and the related 15N- and 13C-TROSY. N-H HSQC measures the environment of the amide N-H bond, and thus provides a single peak for each amino acid in a protein sequence (except for proline; asparagine and glutamine also have primary NH signals). 13C-1H HSQC uses labeled methyl groups to detect changes to valine, methionine, isoleucine, and leucine. If the NMR spectrum has been assigned, changes to the resonances in the presence of compound will suggest the binding site. Even without assigning the protein resonances, however, compounds can be binned by binding site, and non-binders or multi-site binders can be identified. Protein-detected NMR used to be reserved for relatively small proteins; however, technical improvements in NMR hardware and pulse sequences, deuteration of the protein, and selective labeling have made many more proteins amenable to these experiments.

Isothermal calorimetry (ITC): ITC measures the heat generated or absorbed by a binding interaction. For weakly binding PPI inhibitors (in the mid micromolar range), ITC can be challenging because protein usage is high, compound solubility can be limited, and the heats of binding are small. It is important to match the protein and compound buffers and to control for the heat-of-dilution as the compound sample is added to the protein (or vice versa). Despite these challenges, ITC can be very valuable due to the fact that unlike some other methods, ITC is truly label-free, and all components are in solution. By directly measuring the energy of binding, ITC provides information on the entropy and enthalpy of the interaction, the binding affinity (by titrating one of the partners) and the binding stoichiometry. More detail on how to conduct these studies can be found at the MicroCal website.

Thermal stabilization - differential scanning calorimetry (DSC) and differential scanning fluorimetry (DSF, Thermafluor, Protein Thermal Shift): One way to define protein stability is by the temperature at which the protein unfolds. Unfolding is usually a highly cooperative process, and gives a defined melting temperature (Tm) under a given condition of concentration, buffer, etc. When a compound binds to the protein, the complex is more stable than the protein alone, and the protein’s Tm increases. To measure the binding affinity of a compound for a protein, one monitors the increase in Tm (ΔTm) as the concentration of compound is increased. Tm measurements are generally done with micromolar concentrations of protein, and are therefore most sensitive to determining binding affinities in this range. This method also has the advantage that all components are in solution.

There are several methods for measuring the change in Tm. Differential scanning calorimetry (DSC) monitors the heat absorbed by the protein as the temperature is increased; the energy/degree increases at the Tm. Differential scanning fluorimetry (DSF) monitors the binding of a hydrophobic dye to the protein as the temperature is increased. The dye binds preferentially to hydrophobic portions of a protein that are exposed when a protein melts; this binding is accompanied by a change in fluorescence as a function of temperature. Typical DSF dyes include SYPRO orange and 1,8-ANS; DSF measurements are read in specialized instruments or in real-time PCR machines. The magnitude of DSC and DSF signals, and the ΔTms obtained from small-molecule binding studies, is dependent on both the protein and the assay conditions. Thermodynamic statements are only valid in the cases that thermal denaturation is reversible. Nevertheless, ΔTm measurements can provide a rapid assessment of binding affinity, and are increasingly being used in primary screening assays as single-concentration measurements. Sample assay development guidelines can be found in Niesen, et al (23).

Sedimentation Analysis (SA; Analytical ultracentrifugation): Sedimentation analysis measures the sedimentation of proteins in response to a centrifugal force. The protein concentration is measured along the length a centrifugation cell using the proteins absorbance, refractive index, or fluorescence. Two general types of SA experiments are Velocity Sedimentation and Equilibrium Sedimentation. Equilibrium sedimentation gives a first-principle measurement of molecular mass, and is often used to measure self-association (e.g. dimerization) constants. It can also be used, however, to assess the binding of a small molecule to the protein, particularly if the molecule has absorbance at wavelengths distinct from the protein (e.g. > 300 nm). The compound’s aggregation state and the compound’s affect on the apparent molecular mass of the protein provide a quick readout of aggregation-based artifacts. Direct binding of the compound to protein can also be assessed (24). Analytical centrifuges are sold by Beckman Coulter, and add-on fluorescence detection is available from Aviv Biomedical.

X-ray crystallography: X-ray crystallography continues to be the gold standard for characterizing protein/small molecule interactions. The high-resolution (ca. 1.5 – 3 angstrom) structure fit from x-ray diffraction data provides information on the binding site and the specific contacts between compound and protein. The presence of a single molecule bound to a single binding site suggests – but does not prove – that the compound’s inhibition of a PPI arises from binding at that site. Co-structures of compounds and proteins are generally prepared by soaking the compound into a crystal of the protein or by co-crystallization of the protein and compound together. In many cases, it is difficult to obtain co-crystal structures, either because the protein does not crystallize well, the compound induces changes to the protein structure that inhibit crystallization (e.g. binding at a crystal contact, changing the protein conformation), or the compound is not soluble enough.


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Additional References

  1. Degorce F, Card A, Soh S, Trinquet E, Knapik GP, Xie B. HTRF: A technology tailored for drug discovery – a review of theoretical aspects and recent applications. Curr Chem Genomics. 2009;3:22–32. [PMC free article: PMC2802762] [PubMed: 20161833]
  2. Karimova G, Pidoux J, Ullmann A, Ladant D. A bacterial two-hybrid system based on a reconstituted signal transduction pathway. Proc Natl Acad Sci U S A. 1998;95:5752–5756. [PMC free article: PMC20451] [PubMed: 9576956]


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All Assay Guidance Manual content, except where otherwise noted, is licensed under a Creative Commons Attribution-NonCommercial-ShareAlike 3.0 Unported license (CC BY-NC-SA 3.0), which permits copying, distribution, transmission, and adaptation of the work, provided the original work is properly cited and not used for commercial purposes. Any altered, transformed, or adapted form of the work may only be distributed under the same or similar license to this one.

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