Measuring Interactions Between Proteins and Small Molecules or Nucleic Acids

Abstract

Interactions between proteins and small molecules or nucleic acids play a pivotal role in numerous biological processes critical for human health and are fundamental for advancing our understanding of biological systems. Proteins are the workhorses of the cell, executing various functions ranging from catalyzing biochemical reactions to transmitting signals within the body. Small molecules, including drugs and metabolites, can modulate protein activity, thereby impacting cellular processes and disease pathways. Similarly, nucleic acids, such as DNA and RNA, regulate protein synthesis and function through intricate interactions. Understanding these interactions is crucial for drug discovery and development and can shed light on gene regulation, transcriptional control, and RNA processing, providing insights into genetic diseases and developmental disorders. Moreover, studying protein-small molecule and protein-nucleic acid interactions enhances our comprehension of fundamental biological mechanisms. A wide array of methods to study these interactions range in cost, sensitivity, materials usage, throughput, and complexity. Notably in the last decade, new techniques have been developed that enhance our understanding of these interactions. In this review, we aim to summarize the new state-of-the-art methods for detecting interactions between proteins and small molecules or nucleic acids, as well as discuss older methods that still hold value today.

INTRODUCTION:

Understanding how proteins interact with other reagents, specifically nucleic acids, is key to our understanding of many biological functions. For example, examining the interaction between the oncoprotein MDM2 and tumor suppressor p53 revealed the inhibitory function of MDM2 on p53 and opened the door for anti-cancer drugs targeting the MDM2-p53 pathway (Nag, Qin, Srivenugopal, Wang, & Zhang, 2013). Furthermore, elucidating the interactions between proteins and small molecules is critical for successful small molecule-based drug design (Ngo & Garneau-Tsodikova, 2018; Zhong et al., 2021). In the past ten years, many new methods have been developed for examining these interactions. Specifically, fluorescent-based techniques and mass spectrometry have been employed. Many of these techniques offer higher sensitivity, more intricate data, and require much less material. However, these techniques are often expensive and require specialized machinery. As such, more traditional techniques such as competition pulldowns and electromobility shift assays (EMSAs) are still employed. Because of these new advances, the spectrum of how to study the interactions between proteins and small molecules or nucleic acids has become very broad. In this review, we aim to discuss in depth new methods in this field, while also summarizing more traditional methods that still hold value in the field today. Specifically, we will divide these methods into subgroups based on the general type of technique: immobilized binder light-based sensors (IBLBS), solution-based fluorescent techniques (SBFTs), gel-based techniques (GBTs), or mass spectrometry (MS) (Figure 1).

Figure 1:

Breakdown of how the interaction measurements discussed in this review are divided into subgroups (green) and the examples of each that will be discussed (pink). Created with BioRedner.com

Generally, IBLBS immobilize a ligand on a chip or a tip and flow an analyte reporter over the captured ligand, measuring the change in white light flowing through a sensor. SBFTs employ similar techniques, however, both the ligand and reporter are free in solution, and either a labeled dye or labeled analyte is used as a reporter. MS techniques can also measure the interaction between proteins and small molecules or nucleic acids by quantifying mass shifts in the native protein spectra to identify the relationship between a protein and a binding partner. GBTs, in contrast to the previously mentioned techniques, are more traditional assays that rely on the appearance or migration of a complex in a gel matrix to identify an interaction between proteins and various substrates. In this review, we will discuss common methods for each of these techniques, provide the current state of the field, and compare the pros and cons of each.

IMMOBILIZED BINDER LIGHT-BASED SENSORS:

IBLBS utilize the accumulation of a fluorescent signal to quantify the amount and rate of interactions between a protein and a small molecule or nucleic acid. In these techniques, one of the interaction partners, usually called a ligand, is immobilized on either a sensor chip surface or a biosensor tip while the other interaction partner, usually called an analyte, is added in solution (Murali, Rustandi, Zheng, Payne, & Shang, 2022). The immobilization aspect of these techniques allows for a kinetic analysis where there is almost a 1:1 interaction between ligand and analyte and low off-target signaling. However, these techniques also require expensive machinery and engineered reagents specific to the capture mechanism. Technologies such as bio-layer interferometry (BLI) and surface plasmon resonance (SPR) provide real-time label-free (RT-LF) techniques that are effective in monitoring interactions between proteins and small molecules or nucleic acids.

Bio-Layer Interferometry

Bio-layer interferometry (BLI) is a technique that uses white light reflection to measure molecular interactions such as between protein-protein, protein-nucleic acid, and protein-small molecule. What makes this technique stand out among other types of IBLBS techniques is the design of the tip. Immobilized onto these tips is a ligand of interest, usually either a nucleic acid or protein. These ligands are immobilized onto the tips through affinity capture such as the interaction between a Ni-NTA tip and a his-tagged protein, or biochemical capture, such as between a Protein-A coated tip and an antibody (Kumaraswamy & Tobias, 2015). The tip is designed to minimize nonspecific binding by reflecting white light and generating constructive and destructive wavelengths (Kumaraswamy & Tobias, 2015). These wavelengths are then translated into a waveform in real time, revealing when the immobilized ligand interacts with the analyte, in solution by measuring alterations in the biosensor’s opacity and interference patterns (Desai, Di, & Fan, 2019) (Figure 2). These alterations can be measured in real-time to quantify both the association and dissociation between the ligand and analyte, which can be used to calculate binding affinities (Desai et al., 2019). BLI can be utilized to measure even weak interactions between a ligand and an analyte, with a limit of detection lower than 10 pg/mL on most BLI instruments (Mechaly, Cohen, Cohen, & Mazor, 2016). However, one common issue with BLI is that even though the ligand and analyte concentration can be low, the sample volume required is usually high, oftentimes over 400 μL (Desai et al., 2019).

Figure 2:

Diagram of bio-layer interferometry signaling. As molecules bind to the biocompatible surface, the wavelength of the reflected light is altered, causing a shift in the resulting BLI plot. Created with BioRedner.com

When paired with other techniques, BLI can be a powerful tool that determines different protein substrate specificity and binding affinity that would otherwise be difficult to define. A recent study paired cell-free expression (CFE) with BLI to analyze carbohydrate-lectin binding in solution or immobilized on the sensor tip without a purification step (Warfel et al., 2023). A cell-free expression environment allows scientists to evaluate and manipulate protein transcription and translation outside of the cell which can improve the yield and production of low abundant or fragile proteins and protein complexes. Using a streptavidin-coated BLI tip coated with a biotinylated glycan, L-rhamnose, researchers showed they could isolate rhamnose-binding lectins directly from a crude cell-free expression mixture. Lectins are proteins that can recognize and bind certain glycans. Furthermore, they could define the specificity of various Lectins to different types of glycans by evaluating the ability of various saccharides to inhibit the association between the lectins and biotinylated rhamnose using BLI. This in-depth analysis included not only the measure of the amount of binding, but also a determination of binding kinetics such as K_on, K_off, and K_d. Specifically, this competition technique enabled the detection of the lectins binding rhamnose-containing O-antigen through competition with the biotinylated rhamnose on the BLI tip. O-antigen is a Shigella serotype and a targeted molecule in vaccine development (Nowak et al., 2020). Using BLI, researchers were able to validate the use of this technique in measuring the binding kinetics of crude material, as opposed to just purified protein, and also showed translational relevance by the measurement of the interaction between a lectin and O-antigen through competition of the immobilized biotinylated L-rhamnose.

Another study analyzed the protein (rh-Epo)/Multilamellar liposome interactions using streptavidin biosensor tips (Wallner, Lhota, Jeschek, Mader, & Vorauer-Uhl, 2013). The experiment aimed to determine which type of liposomes interact best with rh-Epo. Rh-Epo has neuro-protective properties, where liposomes can be used in vitro to as a model for lipid bilayers. In these assays, researchers aimed to compare the specificity of Rh-Epo to different liposomes to model the preference of Rh-Epo to certain biophysical properties of a membrane. In this experiment, streptavidin BLI tips were loaded with biotinylated rh-Epo protein, placed in a solution with the targeted liposome, and the association was measured in real-time followed by the measure of dissociation in real-time. The results revealed that more saturated lipid chains bind to the protein more readily. Importantly, this assay realized another benefit of BLI over other IBLBs such as surface plasmon resonance (SPR), which will be discussed next in this review. To study the association of weakly associating binding partners, these researchers could evaluate the association phase using BLI for 8000 seconds due to the diffusion-based method for analyte delivery. Comparatively, SPR is often limited to an association phase measurement of roughly 100 seconds due to the need for a microfluidic system to deliver analysts to the sensor surface (Murali et al., 2022). Overall, this study highlighted BLI as an efficient way to determine the most suitable composition of assays for drug discovery and screening research.

Surface Plasmon Resonance

Surface plasmon resonance (SPR) is a method that measures protein-protein, protein-nucleic acid, and protein-small molecule interactions through the reflection of light off a sensor chip bound to a binding agent. Reminiscent of BLI, this method utilizes an immobilized ligand and measures binding as free analytes in a buffer flow across the binding surface. However, unique to SPR, the ligand binding surface is a chip in contrast to a BLI tip. With the sensory chip loaded with an affinity-bound ligand, the binding of the analyte molecules to the ligand is measured in real-time as the refraction index changes according to the mass and charge change caused by the binding of the immobilized molecules (Chung, Park, Bernhardt, & Pyun, 2006) (Figure 3). This technique has evolved exponentially in the past few years with some machines able to measure nearly 400 interactions at a time, putting SPR at the forefront of techniques for high-throughput experiments. This technique is often employed to characterize drug candidates because of its robust and reliable reporting of kinetic data (Matsunaga et al., 2023).

Figure 3:

Diagram of surface plasmon resonance signaling. As more analyte binds, the light reflected from the light source through the prism onto the detector changes in wavelength, causing a shift in the resulting SPR signal. Created with BioRedner.com

SPR, which can be optimized to improve the signal and dynamic range for an individual analyte of interest, has been employed to detect biomarkers, such as cancer biomarkers. For example, a recent study compared the affinity of an anti-hIgG antibody to NeutrAvidin, protein A, NeutrAvidin–protein A complex, and bare gold surface of SPR biosensor as a control (Chung et al., 2006). Once they identified the NeutrAvadin-protein A complex as the most proficient in ligand binding, they were able to further optimize their technique by using self-assembled monolayer (SAM) SPR to improve immobilization. The purpose of this study was to improve the amount of antibody ligand that is bound to the sensor in an optimal orientation for analyte binding. The ability to optimize this step allowed researchers to use less overall antibody reagent and improve the reliability of their binding data. These experiments showed that the NeutrAvidin–protein A complex was most efficient in binding the target molecule at an optimal orientation and that the SAM method further improved their overall signal. After concluding this, the researchers could use this personalized and optimized SPR protocol to detect CEA, a type of cancer marker broadly implicated in a variety of different cancers (Ju et al., 2021). To do this, the anti-CEA antibody and the NeutrAvidin–protein A were added to the surface of the SPR biosensor and the binding of CEA to the immobilized CEA antibody was then measured. Results revealed NeutrAvidin–protein A complex combined with the SAM technique increased the sensitivity to CEA detection and decreased non-specific binding. Thus, SPR can be used to not only determine the binding of molecules to proteins, but it can also be individualized and optimized to target certain specific biomarkers and other structures that would be otherwise difficult to detect.

SOLUTION-BASED FLUORESCENT TECHNIQUES:

SBFTs are unique in the fact they offer quantifiable data about the rate and scale of a protein’s association with small molecules or nucleic acids without the need for immobilization of one of the agents. This usually makes these experiments less expensive and more broadly applicable across a wide range of lab setups. For example, some of these assays can be analyzed in a basic RT-PCR machine or plate reader, which are readily available to most academic or industry labs. The basic principle of these techniques involves the incubation of a ligand and analyte in solution. In this solution, one of the substrates must be fluorescently labeled or a fluorescent dye must be added to quantify the interactions (Horiuchi & Ma, 2009). However, some of these techniques do require a specific machine to visualize the endpoint of these assays and, depending on the assay, require engineered reagents that contain a fluorophore.

Thermal Shift

A molecular biology method called the thermal shift assay combines protein melt temperature analysis with real-time measurement. The central significance of the thermal shift assay is shown through the protein’s melting point analysis. As the temperature changes, the accumulation of signals from sensitive fluorescence dyes is monitored and the thermal denaturation of proteins can be observed (Huynh & Partch, 2015) (Figure 4). The protein denaturation and its stability can be analyzed in different pH, concentrations, sequence mutations, formulations, and temperature conditions using this assay. One common fluorescent dye used in thermal shift is SYBR Green. SYBR green associates with hydrophobic regions of proteins which become exposed upon unfolding caused by the ramping temperature. This dye works well for accurate temperature control and real-time monitoring (Gudnason, Dufva, Bang, & Wolff, 2007). While the binding kinetics that can be gleaned from thermal shift assays are often not as quantifiable as those from SPR or BLI, it does give researchers the ability to understand the functional implications these interactions may have in the protein of interest.

Figure 4:

Thermal shift workflow. Two molecules (i.e., a protein and a small molecule drug candidate) are incubated together and then mixed with a thermal shift dye reagent in a plate where the melting temperature of the protein is then detected using a qPCR machine and constant monitoring of the accumulation of the signal from the dye as the temperature increases. Created with BioRedner.com

One study combined both conventional PCR and thermal shift assay techniques to analyze the effect of protectin DX (PDX) on the expression of the pro-inflammatory cytokines, mediators, and CD14 (Jeon et al., 2024). The examination of thermal shift revealed that PPARγ exhibited increased heat resistance following binding to PDX, suggesting a particular type of interaction between PDX and the PPARγ ligand binding domain. The findings validate the function of PDX leading to anti-inflammatory effects which can be used to treat chronic inflammatory diseases with the involvement of thermal shift RT-qPCR techniques. This discovery broadens the range of methods available for clarifying the relationships that exist between target proteins and bioactive substances within biological environments.

As previously mentioned, thermal shift can be employed to measure how the formulation of a protein (pH, concentration, buffer, etc.) can impact the thermostability of proteins and the interactions between proteins and their binding partners. A group from the University of Braunschweig describes an experiment where the melting point of soluble guanylyl cyclase (sGC), a dimeric heme-containing protein that catalyzed the conversion in GTP to cGMP and plays a role in various diseases, is increased by three known heme-mimetic activator drugs (Elgert, Rühle, Sandner, & Behrends, 2020). In this experiment, the pH, buffer concentration, protein/dye ratio, and protein amount, are manipulated to optimize their sGC sample without the presence of a binding partner. This arm of the study highlighted the utility of thermal shift to examine a protein head-to-head in the presence or absence of a binding partner under various conditions. Furthermore, they were able to confirm the binding of the previously mentioned activators to sGC by observing an increase in the melting temperature of sGC in the presence of the activators compared to the protein alone. This data not only confirmed the association between the activators and sGC but also confirmed that this interaction helped to stabilize the structure of sGC. This data highlights thermal shift as a reliable tool to detecting the association between proteins and small molecules, as well as the ability to analyze the stability of these complexes in different environments and formulations. However, this technique remains only semi-quantitative due to the indirect nature of the measurement as a function of interaction and a moderate limit of detection.

Fluorescence Polarization:

Fluorescence polarization (FP) offers information about molecular interactions within a short timeframe (Lea & Simeonov, 2011). A basic process of fluorescence polarization starts with a beam of light passing through the vertical polarizing filter, which excites the fluorophore, and then the polarized fluorophore can be measured by two conditions. One measures the horizontal light of the excited fluorophore as the perpendicular fluorescence polarization and the other measures the vertical light of the excited fluorophore as the parallel fluorescence polarization (Lea & Simeonov, 2011). The fluorophores’ rotational motion affects the polarization value, which provides data showing the mobility of molecules. When the rotation speed is slow, it will cause a polarized emitted light; on the other hand, when the rotation speed is fast, it will cause an unpolarized or less polarized light emitted (Hall et al., 2016) (Figure 5). FP is most often employed to analyze the association between a larger unlabeled molecule and a smaller fluorescently labeled molecule. For example, when proteins interact with a small, labeled ligand, the rotation speed of the fluorophore will slow and increase the polarization signal. Hence one drawback of FP is that it only gives good and clear information when the size of the labeled protein, ligands, or other molecules is much smaller than that interaction protein, ligands, or other molecules (Dharadhar, Kim, Uckelmann, & Sixma, 2019). However, FP is a useful method for researching different biological processes at the molecular level since it allows researchers to obtain insights into binding affinities, kinetics, and conformational changes. It has shown its value in the drug discovery field of study because it can work with a large group of biological classes that involve secreted proteins, epigenetic regulators, ion channels, other cytosolic proteins, nuclear receptors, kinases, proteases, transcription factors, and other nuclear proteins and enzymes (Hall et al., 2016).

Figure 5:

Diagram of fluorescence polarization signaling. Alone, a small fluorescently labeled molecule has fast rotation leading to a depolarized light emission and low FP signal. When bound to a larger molecule, the labeled small molecule will rotate slower, leading to polarized light emission and a higher FP signal. Created with BioRedner.com

Protein-based fluorescence polarization can be utilized to screen and analyze the activity of SARS-CoV-2 fusion inhibitors by evaluating the binding potency of the SARS-CoV-2 inhibitors, namely HR2 peptide, EK1 peptide, and Salvianolic acid nature product family. The researchers created an expression plasmid with short linkers to connect three HR1 and two HR2 regions of SARS-CoV-2. The expressed protein (the 5-helix bundle construct for SARS-CoV-2) is expected to have a strong binding affinity with SARS-CoV-2 inhibitors, like the mechanism for the 5-HB construct of HIV-1. By using FP, the researchers visualized and compared the interactions between different SARS-CoV-2 inhibitors and the 5-HB construct. FP is a strong tool for investing in molecule interactions and it can set up the fundamental background of further clinical studies (Yin, Chen, Yuan, Liu, & Gao, 2021).

Identifying ciprofloxacin residues in milk that can be used to treat dairy livestock health is directly related to human health through food intake. Researchers from Beirut Arab University presented an FP approach that uses a ciprofloxacin-protein conjugate labeled with a near-infrared fluorescence dye, which we can be used to analyze the change in emission signal that is related to the binding of a monoclonal anti-ciprofloxacin antibody (El Kojok et al., 2020). This assay bypassed the time-consuming incubations and washes and the expensive reagents used in more traditional assays like ELISAs. Without pre-treatment, this quick approach surpasses the EU’s 100 ppb standard with a detection limit of 1 ppb. Experiments that utilize FP save time and money as they use fewer and cheaper reagents compared to the conventional techniques that are laboratory-based, and time is saved because the separation time is not needed based on its single fluorescent label nature.

Protein-based FP can also be used for detecting antibiotic residues in food, which is related directly to human health. The general goal of this study was to design a useful method to detect the concentration of Ciprofloxacin CPFX (antibiotic molecule) in the diluted milk sample. Ciprofloxacin CPFX conjugated with another molecule is labeled with a fluorescence probe (CF647) and utilized to perform the FP assay with a mouse monoclonal anti-ciprofloxacin antibody. Increasing the antibodies will increase the fluorescence signal, based on the principle of FP assay: larger bonded molecules (CPFX with antibody) have higher polarization value; thus, it will have a higher signal. This study also shows the high sensitivity of the FP assay, even if a small amount of CPFX is present, the assay can still detect it: it can detect 1 part of the antibiotic per billion samples based on the FP signals. On the other hand, compared with the other conventional CPFX detection methods (such as liquid chromatography-mass spectrometry), FP is more able to detect CPFX quickly with less specialized instruments. In general, FP is a powerful method for detecting molecular interactions, which not only helps inform developing research but also introduces a new methodology for researchers to determine molecular interactions across many diverse fields in a more accurate and faster way.

Microscale Thermophoresis:

Microscale thermophoresis (MST) is used to study biomolecular interactions between proteins and various ligands (Seidel et al., 2013). This solution-based fluorescence technique avoids issues with surface artifacts and immobilization procedures. An infrared laser is directed into a microscale diameter capillary, where a tiny temperature gradient ranging from 2–6°C is produced and fluorescently tagged molecules in solution are excited by this thermophoretic change (Plach, Grasser, & Schubert, 2017) (Figure 6). Fluorescence is used to track the thermophoretic movement caused by this gradient, which sheds light on the binding of the tagged molecules to target molecules (Romain, Thiroux, Tardy, Quesnel, & Thuru, 2020). Often, the intrinsic fluorescence of tryptophan in proteins or peptides is used for the fluorescent optic tracking system, or extrinsic fluorophores can be used the same way for testing the thermophoretic movement of the molecules. Generally, the intensity of fluorescence in a certain wavelength will change as labeled molecules start to bind to other analytes in solution.

Figure 6:

Diagram of microscale thermophoresis signaling. The laser excites tagged molecules or induces the intrinsic excitement of some amino acids which is then detected as wavelength. Created with BioRedner.com

The use of MST as a method for determining the relative binding affinities between ligands and protein targets is covered in the research from the University of Illinois at Urbana-Champaign (Sparks & Fratti, 2019). They used MST to determine the binding constants for the thermophoretic movement of Sec18 monomer and PA nanodiscs labeled with Atto 647 dye. Their data applied an inverse T-Jump, a thermophoresis analysis method, which was caused by turning off the IR laser after they observed the fluorescence measurement after thermophoretic equilibrium, which only took 10–30 seconds. Furthermore, T-jump, which is caused by opening the IR laser, can happen within 5 seconds, demonstrating this technique’s fast speed and high-quality data.

With minimal sample consumption, the affinity of programmed cell death protein 1 (PD-1) and its ligand, programmed cell death ligand 1 (PD-L1) during tumor escape can be quantitatively analyzed by MST and provides great potential for analyzing the protein-protein interactions in the immune escape research area (Romain et al., 2020) The purpose of this research was to address disagreements in dissociation constant (K_d) values found in other studies by introducing the use of MST for the investigation of the binding interactions between PD-1 and PD-L1. By overexpressing eGFP fusion protein, PD-1, and PD-L1’s affinity constant can be defined without the necessity for any purification steps. This approach reduces the research expense due to the ability of MST to measure the strength of interactions between two molecules with its fluorescence signal detection function under the IR-laser induced temperature variation.

MST offers a wide range of advantages for research. It can offer information about the occurrence of aggregation, the verification of the presence of fluorescence variation through the capillary scans, and be used to determine binding constants for the thermophoretic movement of fluorescent-labeled molecules (Sparks & Fratti, 2019). MST can be used to analyze the protein-protein interactions for immune escape mechanisms (Romain et al., 2020) and provide high-throughput screening with inhibitors to evaluate the ability of specific molecules to modulate the affinity of the target substance in a cheap and fast way.

Time-Resolved Fluorescence Resonance Energy Transfer:

Time-resolved fluorescence resonance energy transfer (TR-FRET) is a sophisticated biophysical technique widely employed in molecular biology and drug discovery. TR-FRET builds upon the foundational principle of Förster resonance energy transfer (FRET), wherein energy is transferred from an excited-state donor fluorophore to a nearby acceptor fluorophore. When the donor fluorophore is excited by an external light source, it can emit energy in the form of photons. If an acceptor fluorophore is within a close vicinity, usually within 10 nanometers, it can capture this energy and subsequently emit light of a longer wavelength (Millar, 1996) (Figure 7). The efficiency of energy transfer is contingent upon the distance and orientation between the donor and acceptor molecules, providing a reliable indicator of molecular interactions. What distinguishes TR-FRET is its integration of time-resolved measurements, allowing precise monitoring of biomolecular interactions over time. By utilizing a time delay between excitation and emission detection, TR-FRET eliminates short-lived background fluorescence, resulting in enhanced signal-to-noise ratios and sensitivity. In practice, TR-FRET assays involve labeling target molecules with donor and acceptor fluorophores, followed by excitation with a pulsed light source and measurement of the resulting fluorescence decay kinetics. This approach enables the quantitative analysis of various molecular processes, including protein-protein interactions, enzyme kinetics, and ligand binding events, with high temporal resolution and accuracy. This proximity-based mechanism renders TR-FRET exceptionally sensitive to subtle changes in molecular conformation or binding events, making it an invaluable tool for studying protein-protein interactions, ligand binding, and structural rearrangements in real time (Millar, 1996).

Figure 7:

Diagram of time-resolved fluorescence resonance energy transfer signaling. When two differentially fluorescently labeled molecules interact, the excitation of one can elicit emission from the other. Created with BioRedner.com

One study researched the ErbB family of proteins, which include four types of receptor tyrosine kinases: Epidermal Growth factor receptor (EGFR), HER2, HER3, and HER4. The three-dimensional structures of the extracellular domain of the ErbB family are known to activate the EGFR signaling pathway. However, modifications to HER can cause malignant transformation of this signaling pathway (Khan, Kryza, Lyons, He, & Hooper, 2021). Therapies that target HER receptors today such as the anti-HER monoclonal antibodies (mAbs), which target the extracellular domain, and HER-specific tyrosine kinase inhibitors (TKIs), which target the tyrosine kinase activity of these receptors are not completely understood and their effects vary from patient to patient (Khan et al., 2021). As a result, this study aims to determine a better biomarker for HER. After evaluating the biochemical approaches commonly used for HER studies and quantitative studies such as co-immunoprecipitation or cross-linking, the researchers determined that TR-FRET is the most sensitive and easiest method to perform when it comes to detecting the dimerization process of native receptors in their biological context (Gaborit et al., 2011). The researchers used terbium cryptate for fluorescence and d2 dye as an acceptor which detects EGFR/HER2 in different cell lines containing different EGFR and HER2 expression levels. The effects were evaluated in an ovarian carcinoma cell line (SKOV-3) and compared with their efficacy in xenografted nude mice. Results revealed that their antibody-based TR-FRET assay was able to detect the formation of homodimers and heterodimers as well as detect the reduction of EGFR/HER2 heterodimers in SKOV-3 cells. The sensitivity of this method was ultimately able to help the researchers take a step closer to understanding the in vivo effects of different target therapies used to suppress tumor growth.

Another cancer study examined the bromodomain (BrD) protein and the histone acetyltransferase (HAT) catalytic domain found in cAMP-responsive element binding protein (CREB) binding protein (CBP) and adenoviral E1A-binding protein (P300) which are targets for therapeutic interventions in cancer and immune system disorders (F. C. Zhang et al., 2020). This study aimed to examine small molecule inhibitors targeting BrD outside of the BrD and extra terminal domain (BET) to develop a novel chemical scaffold for CBP BrD. The researchers developed a TR-FRET HTS assay that targets CBP Brd to analyze and identify small molecules that block the interaction of CBP BrD to H4. The TR-FRET assay developed for this method successfully determined 26 different candidate compounds that inhibit CBP BrD. Since the researchers trusted the well-characterized nature of SPR for hit validation, SPR was then used to confirm the binding affinity and effectiveness in the compounds determined initially as candidates TR-FRET. Results determined that the compounds were successfully bound to CBP BrD thus allowing the researchers to conclude that TR-FRET is a reliable method that can promote drug discovery for CBP BrD interaction. TR-FRET is a robust and high throughput technique for identifying molecular interactions and shows a similar ability to define binding affinity compared to well-characterized and more expensive methods such as SPR.

GEL-BASED TECHNIQUES:

Gel-based techniques (GBTs) rely on the appearance or migration of a band within a gel matrix to quantify the formation of a complex. In general, these more traditional techniques are relatively simple and applicable across a wide range of lab setups. These techniques are also relatively fast and customizable. However, the sensitivity of these assays is usually much lower, requiring microgram amounts of reagents to visualize the data. Furthermore, they often rely on the visualization of a size shift, which makes the interpretation of the binding of small molecules or short strands of nucleic acids difficult to quantify using these techniques. EMSAs employ running an electrophoresis gel in native conditions and comparing the molecular weight of a protein or DNA in the presence of a putative binding partner (Hellman & Fried, 2007). In contrast, competition pulldowns rely on the immobilization of the ligand, usually on a magnetic bead, the binding of the analyte, the addition of free ligand, and the observation of the appearance/disappearance of a band in a gel matrix for the analyte. This technique allows for the identification of associating between a ligand and an analyte, and also for crude quantification of binding kinetics depending on the amount of free ligand added to the solution to cause a decrease in the bound-analyte signal (Louche, Salcedo, & Bigot, 2017).

Electrophoretic Mobility Shift Assay:

Electrophoretic mobility shift assay (EMSA) serves as a pivotal tool in the investigation of protein-nucleic acid interactions due to its sensitivity and versatility. At its core, EMSA involves the mixing of proteins and nucleic acids of interest, followed by their separation via electrophoresis, a process based on the differential migration of molecules primarily dictated by their size and charge. During electrophoresis, unbound DNA migrates through the gel matrix according to its molecular weight, whereas DNA bound to proteins exhibits reduced mobility and appears as distinct, shifted bands. This differential migration pattern allows for the visualization and characterization of protein-DNA complexes (Hellman & Fried, 2007) (Figure 8). Notably, EMSA can accommodate fluorescently labeled components, enabling their detection via various imaging or spectroscopic methods, further enhancing its utility in elucidating molecular interactions. Despite its sensitivity in detecting weak protein-nucleic acid interactions, EMSA may not provide precise quantification for highly complex interactions due to its qualitative nature. Moreover, the technique’s efficacy diminishes when applied to interactions between small molecules, peptides, or pieces of DNA as detection of any shift in size within the gel matrix may be difficult to visualize (Hellman & Fried, 2007). Nonetheless, EMSA remains a valuable tool in the repertoire of molecular biologists for dissecting and characterizing diverse nucleic acid-protein interactions.

Figure 8:

In a western blot against a small piece of DNA, an electrophoretic mobility shift assay will show a band shifted higher in a “DNA + Protein” lane if the two interact with one another. Created with BioRedner.com

The mechanism for NLRP3 inflammasome activation, which has been linked to a multitude of diseases and cancers, remains relatively elusive (Xian & Karin, 2023). In a recent study, a group used EMSA to show for the first time that NLRP3 associates directly with ox-mtDNA and that this interaction may trigger NLRP3-dependent inflammation. In a study that aimed to show that NLRP3 can bind biotinylated non-oxidized and biotinylated oxidized-mtDNA (biot-ox-mtDNA), the group used EMSA to test how ox-DNA is affected by different concentrations of purified NLRP3 by testing the shift of biot-ox-DNA. In this case, EMSA was able to show a shift of the oxidized DNA that was correlated to increasing protein concentration along the same area that NLRP3 migrates on a gel matrix by probing a western blot of the EMSA with a streptavidin antibody against the biotin on the DNA (Cabral, Cabral, Wang, & al, 2023). To test if the ox-DNA shift localizes with NLRP3, the group used a monoclonal NLRP3 antibody to show that the higher concentration of NLRP3 with the antibody was able to match the increase of the shift in ox-DNA. This study illustrated a new function for NLRP3 binding directly to ox-mtDNA and highlighted the ability of EMSA to detect interactions between larger proteins and smaller nucleic acids.

In a recent study that identified a small molecule inhibitor that blocks DNA binding, EMSA was a primary technique utilized that separated unbound ssDNA and a protein-ssDNA complex. Replication protein A (RPA) is a DNA-binding protein found in Trypanosoma brucei, a parasite (Mukherjee et al., 2023). RPA binds ssDNA resulting from damage and recruiting damage repair machinery to the site. They were interested in whether an inhibitor, JC-229, could inhibit the interaction between RPA and ssDNA, thereby inhibiting the repair pathway of this dangerous parasite. In this research study, the group employed EMSA to observe how JC-229 inhibits the ss-DNA binding activity of recombinant RPA proteins (Mukherjee et al., 2023). They were able to show that in the absence of JC-299, ssDNA shifted high in the gel to colocalize with RPA. However, the addition of JC-299 was able to inhibit this shift and associated binding in a dose-dependent manner. Though this data was further bolstered using MST, the initial data gleaned from the EMSA results was a good preliminary confirmation of their mechanism, as well as a good place to optimize their assay without the use of costly MST.

In another study that examined regulating TREM2, a protein associated with Alzheimer’s disease, EMSA was used to confirm their proposed mechanism that shows the interaction between the TREM2 gene and the protein Yin Yang 1 (YY1) (Lu et al., 2023). YY1 is a protein that the group found to be capable of binding the TREM2 gene to regulate the expression of the TREM2 protein. The group shows that EMSA can be used to confirm protein interactions by visualizing a shift in the TREM2 that localized with the YY1 in a dose-dependent manner. The group’s confirmation of YY1 binding the TREM2 gene through EMSA can be further utilized and applied to clinical settings as YY1 can be used to regulate TREM2 expression and help against Alzheimer’s disease.

EMSA is a valuable tool for gleaning preliminary data about protein interactions. While it is a useful tool for assay optimization, EMSA usually must be paired with other techniques as EMSA does not achieve the detailed and quantifiable results of other interaction studies discussed in this review.

Competition Pulldown:

Competition pulldown assays are used to visualize and study protein-protein and protein-DNA. Competition pulldown assays not only show protein interaction but also the binding specificity and strength to a target molecule compared to that of other molecules (Louche et al., 2017). The benefits of competition pull-down assays are that they can validate protein-protein or protein-DNA interactions with the use of an affinity ligand bound to a target/bait molecule that is incubated with another molecule it can interact with. In a pull-down assay, the ligand of interest with an affinity tag is immobilized on a magnetic bead, then it is incubated in the analyte proteins, oftentimes over a concentration gradient (Louche et al., 2017) (Figure 9). The nonspecifically bound analyte is washed away, and the specifically bound analyte can be eluted from the immobilized ligand by adding a free ligand to the complex, which can also be done over a concentration gradient. The results are usually measured by western blot against the analyte. Pull-down assays, can, however, yield false positives and negatives if, for instance, the washes are not stringent enough to get rid of non-specifically bound proteins, and they often rely on being paired with other techniques to get complete and quantifiable results.

Figure 9:

Competition pulldown workflow. Protein is bound to biotinylated DNA immobilized on streptavidin beads, then competed off the beads with varying amounts of unlabeled DNA to show DNA binding, specificity, and dose dependency. Created with BioRedner.com

In a study focused on the differential binding of NLRP3 to ox-mtDNA and non-oxidized-mtDNA to mediate the NLRP3 inflammasome, a pull-down assay was used to show that NLRP3 can differentially bind oxidized and non-oxidized DNA. The magnetic streptavidin beads used were incubated with biotinylated oxidized or non-oxidized mitochondrial DNA and the unbound DNA was washed away (Cabral, Cabral, Wang, Zhang, et al., 2023). NLRP3 protein was then added to the beads to bind to the immobilized DNA, and then it was eluted using increasing concentrations of oxidized or non-oxidized mtDNA. As the concentrations used increased, the eluted fraction of NLRP3 also increased, indicating binding. A western blot against NLRP3 confirms that NLRP3 differentially binds oxidized and non-oxidized DNA. Furthermore, the NLRP3 construct was a truncated mutant containing only the first domain of the three-domain full-length NLRP3: the pyrin domain. Thus, the pulldown assay offered crucial insight into the pyrin domain’s specificity towards ox-mtDNA due to its higher affinity compared to non-oxidized DNA. This study shows that pull-down assays can be both utilized and modified to see, confirm, and study protein-DNA interactions.

MASS SPECTROMETRY:

Mass spectrometry (MS) is a broadly used technique to classify the identity of a variety of heterogeneous complexes and even identify the sequence of unknown proteins. Specific techniques may be employed using MS to study the interactions between proteins and small molecules or nucleic acids. Intact mass or native MS are unique tools that report the mass size of undigested protein samples. By leaving the protein intact, one may analyze the addition of a small molecule or nucleic acid on the protein as a mass shift (Bennett, Nguyen, & Donald, 2022). Not only is this technique sensitive enough to sense small molecules or near single nucleotide additions, but it also offers the statistic of the size of the new mass, allowing researchers to conclude the identity of the addition (if unknown) or if there are any higher order species formed through this interaction such as multimers. Stable isotope labeling by amino acids in cell culture (SILAC) also offers the ability to study the addition of small molecules to proteins in a cellular context and has specifically been utilized to identify small-molecule drug targets in cells (Ong, Li, Schenone, Schreiber, & Carr, 2012).

Intact Mass

Traditional applications of mass spectrometry utilize fragmentation to detect mass shifts in a protein construct. Alternatively, intact mass spectrometry (intact MS) utilizes the undigested native conditions of a protein, enabling quantitative analysis of its structure and non-covalent interactions with other proteins and/or DNA (Hale, Illes-Toth, Mize, & Cooper, 2020). Once purified, a protein sample is charged through one of two common ionization techniques: Electrospray Ionization (ESI) or Matrix-Assisted Laser Desorption/Ionization (MALDI). In ESI, the protein of interest is dissolved and sprayed through a charged capillary tube using high voltage (Claydon, Davey, Edwards-Jones, & Gordon, 1996). ESI can be advantageous due to its compatibility with a wide range of molecular weights, soft ionization for more fragile constructs, and its ability to be coupled with liquid chromatography. In MALDI, the protein of interest is combined and co-crystallized with a respective matrix compound on a target plate. The protein sample is then ionized through laser irradiation (Claydon et al., 1996). This method can be beneficial for low-expression and fragile protein constructs that are prone to fragmentation. MALDI also provides more precise mass measurements, especially for higher-order species, in which larger-scale post-translational modifications can be maintained and analyzed accurately based on interactions with small molecules or nucleic acids. Upon ionization, the undigested protein sample is measured on a mass spectrometer. The spectra reveal the intensity of ions at specific mass-to-charge ratio values (Figure 10). Preliminary conclusions can be made based on a detectable mass shift concerning the undigested protein alone. The main advantage of analyzing a non-denatured protein complex is the preservation of these post-translational modifications. For example, detectable changes in conformation or binding stoichiometry can provide insight into the functionality of the complex when associated with other small molecules and/or nucleic acid sequences.

Figure 10:

Example of an intact mass plot. In this example, a monomeric protein (left) binds to a small peptide (middle) which induces the dimerization of the larger protein (right). Created with BioRedner.com

Recent applications have made intact MS a useful technique in routine drug screening. For example, researchers employed intact MS to assess the efficacy of meropenem, a carbapenem β-lactam antibiotic, in inhibiting a transpeptidase (LdtMt2), crucial in the formation of the bacterial cell wall in mycobacterial species such as Mycobacterium tuberculosis (Zandi & Townsend, 2021). An analysis with the intact protein displayed instability in the Ldt-meropenem adduct, characterized by decarboxylation and its off-loading by faropenum before prolonged incubation, thus revealing the inefficacy of meropenem against this class of transpeptidases. A reversible reaction and an off-loading reaction that occur within the Ldt-meropenum adduct are measured via detectable mass shifts on the spectra – further highlighting the importance of intact MS, specifically in providing insight into protein-small molecule interactions.

Stable Isotope Labeling by Amino Acids in Cell Culture

Stable isotope labeling by amino acids in cell culture (SILAC) utilizes MS to make accurate proteomic quantifications. These quantifications can be used to characterize stable interactors with proteins, such as small-molecule drug targets. Although proteomic quantifications are crucial in analyzing activation and signaling cascades in various protein complexes, such as detecting phosphorylation events and their association with the activation of a specific protein complex, SILAC is not limited to protein-small molecule interactions, as post-translational modifications can also be detected in protein-DNA interactions (G. Zhang & Neubert, 2009). In SILAC, cells are cultured in a medium that is supplemented with amino acids that contain a specific light (standard) or heavy stable isotope. The cells metabolically integrate the labeled amino acids into newly synthesized proteins (Mittler, Butter, & Mann, 2009). Differing from intact MS, SILAC medium is supplemented with a dialyzed serum, flushing out non-labeled amino acids, including small molecules necessary for cell multiplication. Cells are then harvested, and the proteins are extracted. The isotope-labeled peptides are analyzed by a mass spectrometer, yielding identifiable mass shifts and relative peptide abundance due to the differential labeling and the peak ratios produced from the light and heavy isotopes (Figure 11). While SILAC can detect post-translational modifications in protein-nucleic acid interactions, its main purpose is to provide insight into protein-small molecule interactions, notably small molecule drug targets in the clinical context. Cells isotopically labeled with SILAC and treated with potential small molecule drug targets with specific functionality respective to the protein complex can be analyzed for abundance and/or post-translational modifications using MS, giving useful insight into their association (Spruijt, Baymaz, & Vermeulen, 2013). For example, a potential small molecule activator when combined in a SILAC medium with a specific protein complex, should theoretically show increased abundance via mass shifts.

Figure 11:

Example of stable isotope labeling by amino acids in cell culture plot. Mass shifts related to the heavy/light integrated proteins from the SILAC culture can inform how protein expression and functionality may change in response to an interaction with another protein, piece of DNA, or small molecule. Created with BioRedner.com

Recent applications of SILAC have enabled researchers to confirm novel regulators of inflammasome activation, specifically the association between the phosphorylation of Bruton’s tyrosine kinase (BTK) and the NOD-like receptor family, pyrin domain-containing 3 (NLRP3) protein complex activation (Liu et al., 2017). In this study, researchers used SILAC medium stimulated with varying concentrations of an NLRP3-activator, nigericin. Upon 5 minutes of nigericin addition, BTK regulatory site tyrosine 551 (Y551) displayed rapid phosphorylation and considerable upregulation (2.6-fold in 5 minutes). The phosphorylation and upregulation of the Y551 residue by nigericin + SILAC medium is conclusive that the interactions coincide with NLRP3 inflammasome activation, which shares the same stimulator. A future assay can employ the use of known small-molecule activators/inhibitors to the SILAC medium, afterward analyzing the relative protein abundance and occurrence of phosphorylation and inflammasome activation. Not only does this provide structural insight into the protein construct, but applications in the clinical realm as well.

CONCLUDING REMARKS:

Amongst the diverse array of techniques available for evaluating interactions between proteins and other biomolecules, each offers distinct advantages and limitations. We have summarized each technique and its pros and cons herein (Table 1). In this review, we have divided the techniques into sub-categories: immobilized binder light-based sensors (IBLBS) solution-based fluorescent techniques (SBFTs), gel-based techniques (GBTs), and mass spectrometry (mass spec). For IBLBS, we discussed the use of bio-layer interferometry (BLI) and surface plasma resonance (SPR). BLI and SPR both provide real-time kinetic data with minimal sample preparation, but BLI specifically may suffer from non-specific binding, and SPR requires considerable optimization. The SBFTs discussed included thermal shift, fluorescence polarization (FP), microscale thermophoresis (MST) and time-resolved fluorescence resonance energy transfer (TR-FRET). The most traditional assay of the four, thermal shift, offers simplicity and can analyze ligand binding but may lack specificity. FP, MST, and TR-FRET are all more sensitive and quantifiable. FP assays offer high specificity but require purified components and may not be suitable for large-scale studies, MST offers exceptional sensitivity and can analyze low-affinity interactions but may be limited by sample quality, and TR-FRET offers high spatial resolution and can monitor dynamic interactions in real-time but requires careful optimization and controls. The gel-based techniques of EMSAs and competition pulldowns both offer simplicity and scalability but may lack sensitivity, are often difficult to quantify, and often require purified components. Finally, we discussed the mass spectrometry-based techniques of intact mass and SILAC. Intact Mass analysis provides precise molecular weight information but requires specialized instrumentation, while SILAC enables quantitative analysis of protein interactions but may be challenging for some applications where the cellular implications of these interactions are still unknown. By understanding the strengths and weaknesses of each method, researchers can select the most appropriate approach for their specific experimental needs, advancing our understanding of complex molecular interactions in biological systems.

Table 1.

Overview of techniques used to study the interaction between proteins and small molecules or nucleic acids

Technique	Type of technique	Overview	Common use	Pros	Cons
BLI	IBLBS	A label-free analytical technique that measures biomolecular interactions in real-time as biomolecules bind to a sensor surface.	Analysis of protein-protein, protein-nucleic acid, and protein-small molecule interactions. Real-time monitoring capability, enabling dynamic insights into binding kinetics.	High in sensitivity and can detects weak interactions.	Bulk sensing technique can lead to challenges in distinguishing specific interactions from nonspecific background signals. Require relatively larger sample volumes.
Competition pulldowns	GBT	Visualizes protein-protein interaction and binding specificity of a protein.	protein-small molecule	They can verify protein-protein interactions and map out binding domains on the target protein.	Can yield false negatives and are semi-quantitative so additional methods are needed for better results.
EMSA	GBT	Can detect protein complexes with nucleic acids.	Both protein-small molecule and protein-NA	Very sensitive so they can detect weak- Protein-DNA interactions.	Sensitive so it cannot provide good resolutions for mixtures that are complex.
FP	SBFT	A method for measuring the degree of alignment of fluorescent molecules as they release light after excitation is called fluorescence polarization.	Both highly mobilized and less mobilized molecules.	Important details regarding the molecular environment and interactions in the solution can be learned from the degree of polarization.	The fluorescence polarization approach has certain limitations. Firstly, it can only be applied to molecules that are easily tagged with fluorophores due to their dependence on fluorescent molecules.
Intact Mass	MS	Analyzes mass shifts within an intact protein construct prior to enzymatic digestion or fragmentation.	Both protein-small molecule and protein-amino acid.	Provides comprehensive analysis for protein-small molecule and nucleic acids.	Any modifications made to the construct post-digestion will not be assessed.
MST	SBFT	A biophysical method used to study biomolecular interactions focuses on the thermodynamics and binding affinities of molecules such as ligands, proteins, and nucleic acids.	Ligands, Nucleic acids, and proteins	The MST technique is a flexible and effective tool for studying molecular interactions because of its many benefits, which include its wide dynamic range, low sample consumption, high sensitivity, and compatibility with a variety of biomolecules.	The sensitivity of the MST technology to sample circumstances and attributes may be a possible restriction, affecting measurement accuracy and repeatability.
SPR	IBLBS	Studies molecular interactions in real-time using light coupled with electrons oscillated from metal and dielectric.	Analysis of protein-protein, protein-nucleic acid, and protein-small molecule interactions.	More affordable and consumes less amounts of samples. Provides real-time molecular interactions.	May require prior cloning, such as mutational analysis which can be challenging due to its time-consuming nature. Less sensitive than BLI.
SILAC	MS	A quantitative proteomics assay that utilizes isotopically labeled amino acids to assess relative protein abundance and mass differences.	Protein-small molecule	Labeling sensitivity provides accurate yield.	Cost availability and limited to protein quantifications only.
Thermal Shift	SBFT	It provides the sensitive detection and quantification of RNA molecules and protein by detecting variations in melting temperature during PCR amplification.	Molecules and proteins	Temperature changes in the reaction mixture are observed while fluorescence is tracked, providing information on the melting point and stability of DNA.	Specific thermal cyclers with fluorescence detection capabilities are needed.
TR-FRET	SBFT	spectroscopic technique used to study molecular interactions by measuring the transfer of energy between two fluorophores	Proteins and Nucleic Acids	It does not depend on the excitation intensity or the duration of light exposure and is therefore extracted from the decay of fluorescence intensity.	This technique is very sensitive to the quality and purity of the samples, and experimental conditions need to be carefully controlled to ensure accurate and reliable results.