Expanding the Reach of Membrane Protein-Ligand Interaction Studies through the integration of Mass Spectrometry and Membrane Mimetics

Expanding the Reach of Membrane Protein-Ligand Interaction Studies through the integration of Mass Spectrometry and Membrane Mimetics | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL PROTEOMICS This is a preprint and has not been peer reviewed. Data may be preliminary. 13 August 2025 V1 Latest version Share on Expanding the Reach of Membrane Protein-Ligand Interaction Studies through the integration of Mass Spectrometry and Membrane Mimetics Authors : Jonathon Lambos , Ashim Bhattacharya , Mohammed Al-Seragi , and Franck Duong 0000-0001-7328-6124 [email protected] Authors Info & Affiliations https://doi.org/10.22541/au.175508838.89786308/v1 Published PROTEOMICS Version of record Peer review timeline 685 views 369 downloads Contents Abstract Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Mass spectrometry (MS) offers robust, label-free approaches for characterizing ligand-protein interactions through two main strategies: affinity-based and stability-based assays. Affinity-based methods, such as affinity-selection MS (AS-MS) and mass spectrometry binding assays (MSBA), detect ligand-target interactions by identifying bound ligands or measuring displacement of a reporter ligand. Stability-based techniques, including thermal proteome profiling (TPP) and limited proteolysis-MS (LiP-MS), infer interactions based on changes in a protein’s thermal or proteolytic stability. These MS-based workflows enable proteome-wide analyses with high specificity and throughput. However, their application to membrane proteins (MPs)—a major class of drug targets—has been limited by challenges such as structural complexity, low native expression, and poor compatibility with detergent-based MS protocols. Recent advances in membrane mimetic (MM) systems, including nanodiscs, Peptidiscs, and styrene-maleic acid (SMA) polymers, help address these barriers by maintaining native-like lipid environments and preserving functional MP conformations. These mimetics facilitate proteome-scale solubilization of MPs in forms compatible with MS screening. This review outlines key affinity- and stability-based MS approaches and examines their adaptation for MPs. It also highlights how combining MS techniques with MM systems is expanding the reach of high-resolution, functional analysis of MP–ligand interactions. Expanding the Reach of Membrane Protein-Ligand Interaction Studies Through the Integration of Mass Spectrometry and Membrane Mimetics Jonathon C. Lambos, Ashim Bhattacharya, Mohammed Al-Seragi, Franck Duong van Hoa* Department of Biochemistry and Molecular Biology, Faculty of Medicine, Life Sciences Institute, University of British Columbia, Vancouver, British Columbia, Canada, V6T 1Z3. *Correspondence to [email protected] List of Abbreviations 1. Mass Spectrometry (MS) 2. Affinity-Selection Mass Spectrometry (AS-MS) 3. Mass Spectrometry Binding Assays (MSBA) 4. Thermal Proteome Profiling (TPP) 5. Limited Proteolysis-Mass Spectrometry (LiP-MS) 6. Membrane Protein (MP) 7. Styrene Maleic Acid (SMA) 8. Liquid Chromatography–Tandem Mass Spectrometry (LC-MS/MS) 9. Stability of Proteins From Rates of Oxidation (SPROX) 10. Membrane Mimetic (MM) 11. Membrane Scaffold Protein (MSP) 12. Size Exclusion Chromatography (SEC) 13. Dodecyl Beta Maltoside (DDM) 14. GLP-1 Receptor (GLP-1R) 15. Nicotinic Acetylcholine Receptor (nAChR) 16. Adenosine Triphosphate (ATP) 17. Cellular Thermal Shift Assay (CETSA) 18. Enzyme-Linked Ammunosorbent Assay (ELISA) 19. Differential Scanning Fluorimetry (DSF) 20. Melting Temperature (T m ) 21. Half Maximal Inhibitory Concentration (IC₅₀) 22. Dissociation Constant (K d ) 23. Dissociation Rate Constant (K off ) 24. Lauryl Maltose Neopentyl Glycol (LMNG) 25. Tandem Mass Tag (TMT) 26. Cell-Surface Thermal Proteome Profiling (CS-TPP) 27. Tandem Mass Tag-Based Data-Dependent Acquisition (TMT-DDA) 28. Data-Independent Acquisition (DIA) 29. Isothermal Shift Assay (iTSA) 30. Single-Tube Thermal Proteome Profiling With Uniform Progression (STPP-UP) 31. Drug Affinity Responsive Target Stability (DARTS) 32. Proteome-Wide Elution Profile Learning for Small-molecule Activity (PELSA) 33. Thermostability-Assisted Limited Proteolysis Mass Spectrometry (TALiP-MS) 34. Styrene Maleic Acid Lipid Particle (SMALP) 35. Calcium-Sensing Receptor (CaSR) 36. β₂-adrenergic receptor (β₂-AR) 37. Scintillation Proximity Assay (SPA) 38. Amyloid-β Oligomers (AβOs) 39. Membrane Mimetic Thermal Proteome Profiling (MM-TPP) 40. Cross-Linking Mass Spectrometry (XL-MS) Keywords 1. Mass Spectrometry 2. Protein-Ligand Interaction 3. Membrane Proteins 4. Membrane Mimetics 5. Screening Word Count (excluding references and figures/table legends): Abstract Mass spectrometry (MS) offers robust, label-free approaches for characterizing ligand-protein interactions through two main strategies: affinity-based and stability-based assays. Affinity-based methods, such as affinity-selection MS (AS-MS) and mass spectrometry binding assays (MSBA), detect ligand-target interactions by identifying bound ligands or measuring displacement of a reporter ligand. Stability-based techniques, including thermal proteome profiling (TPP) and limited proteolysis-MS (LiP-MS), infer interactions based on changes in a protein’s thermal or proteolytic stability. These MS-based workflows enable proteome-wide analyses with high specificity and throughput. However, their application to membrane proteins (MPs)—a major class of drug targets—has been limited by challenges such as structural complexity, low native expression, and poor compatibility with detergent-based MS protocols. Recent advances in membrane mimetic (MM) systems, including nanodiscs, Peptidiscs, and styrene-maleic acid (SMA) polymers, help address these barriers by maintaining native-like lipid environments and preserving functional MP conformations. These mimetics facilitate proteome-scale solubilization of MPs in forms compatible with MS screening. This review outlines key affinity- and stability-based MS approaches and examines their adaptation for MPs. It also highlights how combining MS techniques with MM systems is expanding the reach of high-resolution, functional analysis of MP–ligand interactions. Introduction Mass spectrometry (MS), long regarded as the gold standard for protein identification, has evolved into a powerful tool for mapping protein–ligand interactions. With the advent of label-free detection formats, MS now occupies a critical intersection between structural proteomics and drug discovery. This evolution did not occur abruptly but unfolded gradually, paralleling a broader shift in proteomics from merely cataloguing proteins to probing their functional roles within the cellular environment. [1] In protein-ligand studies, this transition toward functional interrogation has been driven by two main classes of MS-based methodologies: affinity-based and stability-based approaches, named according to their underlying mechanisms. [2] Affinity-based techniques, such as AS-MS and MSBAs, directly capture protein–ligand complexes, which are then analyzed via liquid chromatography–tandem mass spectrometry (LC-MS/MS) to reveal bound ligands. [3, 4] This workflow, despite being technically challenging, preserves native interactions and provides straightforward readouts. Modernization in high-throughput formats has dramatically scaled these assays, with pooled AS-MS screens now encompassing libraries of over 20,000 compounds. [5] Stability-based approaches, by contrast, infer binding events indirectly through changes in protein conformation. Methods such as stability of proteins from rates of oxidation (SPROX), TPP, and LiP-MS detect ligand engagement by measuring thermal shifts, oxidation rates, or altered proteolytic cleavage patterns. [6-8] These methods ask not “what binds,” but “is binding altering the protein?” In other words, thermal shifts or altered cleavage patterns serve as proxies for ligand engagement. [9] TPP, in particular, has proven effective for profiling both on- and off-target interactions in whole-cell lysates, offering systems-level insights into compound activity. [10-14] Despite these advances, MPs, which represent nearly 70% of the druggable proteome, remain underrepresented in high-throughput MS screening campaigns. [15, 16] This exclusion is due to a set of enduring biochemical challenges: MPs are often present in low abundance, structurally complex, and embedded in lipid bilayers that resist solubilization. Detergents, though essential for MP extraction from the lipid bilayer, can disrupt native conformations, destabilize ligand-binding sites, or interfere with downstream MS analysis. [16, 17] The narrow window between effective solubilization, preservation of native structure, and compatibility with other assays makes meaningful MP–ligand interrogation particularly difficult. [18] Nonetheless, emerging studies suggest that with suitable adaptations, MS-based methods can be extended to MPs, reinforcing the need for continued innovation in this area [10, 19-21]. As part of these innovations, researchers have begun exploring the use of membrane mimetics (MMs), engineered scaffolds that stabilize MPs in aqueous environments while preserving as much as possible their native conformations. These tools not only support protein stability but also make MPs more amenable to diverse analytical workflows. Among these, nanodiscs, SMA copolymers, and the Peptidisc have been adopted with success. Nanodiscs create a bilayer-like environment by reconstituting MPs into disc-shaped structures, although their preparation often demands significant quantities of purified proteins and the addition of exogenous lipids. [22] SMA copolymers, on the other hand, extract MPs directly from native membranes, offering a native-like environment that preserves endogenous lipids; however, the polymer can introduce chemical interferences during MS analysis. [23] Finally, the Peptidisc, stabilizes MPs post-detergent extraction by reconstituting them into a flexible, multicopy and lipid-free peptide scaffold. The system is characterized by its “one-size-fits-all” design, offering broad compatibility with a variety of MPs and demonstrated simplicity for integration with proteome-wide MS workflows. However, the presence large amount of the Peptidisc scaffold during MS analysis can hinder the detection of low-abundance MPs. [24] As MS-based ligand discovery workflows continue to scale in sensitivity and throughput, the integration of MMs offers not only a technical workaround but a conceptual framework for re-engaging a class of proteins long sidelined by solubility barriers. Together, it is expected that these continuously evolving technologies will offer a promising conduit for integrating MPs into modern drug discovery pipelines. This review, therefore, aims to address two fundamental questions: (i) what are affinity- and stability-based MS approaches, and how have they been leveraged to identify MP–ligand interactions? and (ii), what are the challenges with the identification of MP-ligand interactions, and how do MMs help to overcome these obstacles? 2. Affinity Based Methods 2.1 Affinity Selection Mass Spectrometry Affinity Selection Mass Spectrometry (AS-MS) exemplifies the core strength of affinity-based screening that is, capturing protein–ligand complexes in solution and identifying bound molecules directly by mass, all without requiring chemical modification or labeling. [3] This approach has become increasingly valuable in early-stage drug discovery, where both analytical clarity and throughput are essential. [5, 19, 25-27] In a typical AS-MS workflow, a purified protein of interest is immobilized, often via an epitope tag, on a solid support such as magnetic or agarose beads ( Fig. 1A ). After incubation with a ligand mixture, unbound and weakly bound ligands are removed using methods like vacuum filtration, ultrafiltration, or size exclusion chromatography (SEC). The remaining ligands are eluted with denaturants (e.g. organic solvents or detergents) and identified via LC-MS/MS, with mass-to-charge ratios and retention times serving as key ligand identifiers [5, 25-27]. This method enables screening of complex mixtures, such as compound libraries or natural product extracts, without the need for ligand labeling nor prior purification. [3, 25] AS-MS performs with high sensitivity and precision when applied to soluble proteins. However, its extension to MPs presents significant biochemical and technical challenges—not due to the method itself, but due to the instability and heterogeneity of MPs when removed from their native lipid environments. For many multipass MPs (e.g., GPCRs), detergent purification often induces structural distortion or conformational bias, which directly impacts ligand detection. Nonetheless, adaptations of AS-MS to detergent-solubilized MPs have shown promise. For example, Zhang et al. (2020) used AS-MS on dodecyl beta maltoside (DDM)-purified 5-HT₂C receptors immobilized on magnetic beads, identifying aporphine alkaloid compound 1857, a G protein-biased agonist that selectively activates 5-HT₂C over 5-HT₂A and 5-HT₂B, with implications for weight loss. [27] Despite the potential, other studies have reported a tendency for ligand discovery to favor antagonists over functionally relevant modulators. The risk of conformational bias was illustrated by Lu et al. (2019), who screened a thermostabilized A₂A adenosine receptor against 1,100 compounds. They identified Fg754, a negative allosteric modulator. However, their use of a thermostabilized apo A₂AR variant, initially designed for crystallization studies, likely restricted the interaction of ligands that bind the receptor in different conformations. [5] Thermostabilization, while beneficial for expression and handling, often limits receptor dynamics that are critical for engaging certain classes of ligands. Recognizing the limitation of conformation bias, researchers have shifted toward screening in native membrane environments. Qin et al. (2018) developed an AS-MS workflow using crude insect cell membranes containing GLP-1 receptors (GLP-1R) as bait. This setup preserved native conformation and enabled in situ ligand binding, followed by filtration and LC-MS identification. The study isolated 18 ligands, including four novel positive allosteric modulators. Notably, these ligands failed to bind a thermostabilized GLP-1R mutant, underscoring the importance of preserving native MP dynamics during screening; dynamics that detergent purification often disrupts. [26, 28] While such advances are encouraging, AS-MS faces inherent limitations that constrain its broader application to MPs. First, detection sensitivity depends heavily on the MS platform, and ligands with poor or inconsistent ionization, even if tightly bound, may go undetected [30]. Additionally, protein-ligand complexes can dissociate during washing or separation steps. Protocol optimizations (e.g., adjusting protein concentration, ligand ratios, buffer conditions) can reduce these losses, but do not fully resolve the challenges unique to MPs in detergent systems. [5, 26, 29] The challenge extends to input requirements as well. AS-MS often demands substantial quantities of purified protein, which poses a logistical bottleneck for MPs due to their low expression yields. [29, 30] This alone contributes to the underrepresentation of MP targets in AS-MS campaigns, despite their relevance in pharmacology. 2.2 Mass Spectrometry Binding Assays Unlike AS-MS, which directly detects ligand binding, Mass Spectrometry Binding Assays (MSBA) is a competitive technique that utilizes a non-labeled, MS-detectable reporter ligand, referred to as a marker. Rather than measuring the presence of bound ligands, the primary readout in MSBA is the extent to which test compounds displace the pre-bound marker from its binding site. [29-31] In a typical MSBA workflow, a compound library is divided into sub-libraries and screened in parallel under two conditions: one to measure total marker binding, and the other including an excess of unlabeled competitors to determine specific binding ( Fig. 1B ). Following incubation, unbound compounds are removed through a washing step, and the remaining bound ligands are eluted and identified by LC-MS/MS. [29, 32] This washing step is a critical assay parameter and largely depends on the affinity of the reporter ligand. High-affinity markers allow for stringent washing via filtration, while weaker interactions may require gentler separation methods such as density-gradient centrifugation. Early studies demonstrated that MSBA could be conducted in membrane preparations without detergent solubilization. For example, Sichler and Hofner (2018) employed filtration using a deuterium-labeled reporter ligand, [²H₆]MB327, to investigate nicotinic acetylcholine receptor (nAChR) interactions and displacement by unlabeled competitors. In this study, nAChRs were presented in their native conformation using membrane preparations from Torpedo californica electroplaque tissue, allowing quantification of specific binding in a physiologically relevant context. [33] In a related effort, Chen et al. (2017) expressed the human BLT1 receptor in HEK293 cells and developed the MSBA workflow using membrane preparations. They characterized two reporter ligands with high affinity and strong MS signals, ensuring precise competition-based measurements. Their assays provided kinetic parameters, including ligand–receptor dissociation constants (K d ) and dissociation rate constant (K off ), and confirmed stable ligand retention during wash steps. [34] These controls not only validated the method’s reliability but also underscored that MSBAs can function effectively without labeled ligands. Gabriel and Wanner (2019) subsequently expanded the approach by creating a two-step workflow that integrates MSBA with AS-MS sequentially. Using the GAT1 neurotransmitter transporter as a model, they first employed marker-based competition assays to identify potential binders from a compound library. These candidates were then validated via AS-MS to confirm direct interactions. [30] This hybrid strategy harnesses the strengths of both methods: MSBA offers insight into relative binding affinities and kinetics, while AS-MS provides structural confirmation of ligand identity. However, MSBA is not without limitations. The method depends fundamentally on the availability of a well-behaved marker ligand, one with high specificity, favourable ionization and stable binding. Without such a reference, competitive displacement cannot be measured. [29, 35] Additional complications arise in membrane-rich preparations, where non-specific hydrophobic interactions between lipophilic compounds and bilayer components can elevate background signal and reduce assay fidelity. [28] These effects are particularly pronounced when working with native membranes or crude extracts, where off-target partitioning may obscure true displacement. Despite these challenges, MSBA remains a powerful yet underutilized tool for probing ligand interactions in near-native systems. As interest in membrane-targeted therapeutics grows, refining the biochemical and analytical parameters of MSBA workflows will be critical. Advances in mimetic systems, which preserve MP topology while reducing nonspecific background binding, may play a central role in extending this technique to a wider range of MP targets. 3. Stability Based Methods 3.1 Stability of Proteins from Rates of Oxidation Stability of Proteins from Rates of Oxidation (SPROX) is a chemically-based strategy for detecting ligand-induced changes in protein stability. It assesses how ligand binding alters the unfolding profile of a protein under denaturing conditions. In that case, SPROX employs methionine oxidation as a structural probe: as proteins unfold in the presence of chemical denaturants (e.g., urea or guanidine hydrochloride), previously buried methionine residues become exposed to hydrogen peroxide, leading to their oxidation to methionine sulfoxide, a +16 Da mass shift readily detected via MS. [36, 37] Following denaturation and oxidation, protein samples are enzymatically digested and peptides analyzed by LC–MS/MS. Changes in methionine oxidation across a gradient of denaturant concentrations yield a measurable profile of protein folding stability. Ligand binding that stabilizes the protein shifts methionine exposure to higher denaturant concentrations, indicating delayed unfolding. Conversely, ligand binding may induce destabilization, accelerating unfolding, and shifting oxidation to lower denaturant levels. These spectral shifts, interpreted across conditions or time points, reflect the net energetic impact of ligand binding ( Fig. 2A) . Although SPROX was initially developed for soluble proteins, it has also shown promise for membrane-associated targets. For example, Geer et al. (2016) applied SPROX to map the adenosine triphosphate (ATP) interactome in Saccharomyces cerevisiae , identifying 28 ATP-sensitive proteins, including membrane-associated targets such as the plasma membrane H⁺-ATPase (PMA1) and the V-ATPase subunit VMA1. [20] By combining SPROX with isobaric labeling, the study enabled multiplexed quantification within a single MS run, highlighting SPROX’s potential for interrogating MPs in complex lysates. Notably, both stabilization and destabilization effects were observed, indicating that SPROX can report not only direct ligand interactions but also allosteric effects and broader conformational changes. However, the method has important limitations. Most notably, its dependence on methionine-containing peptides, which restricts coverage in proteins or domains where methionine residues are sparse, structurally buried, or sequestered within the membrane bilayer. [38] This limitation is especially pronounced in transmembrane helices and lipid-embedded sequences, and as a result, certain classes of MPs may be underrepresented or entirely missed in SPROX datasets. 3.2 Thermal Stability Methods 3.2.1 The Cellular Thermal Shift Assay The Cellular Thermal Shift Assay (CETSA) leverages a fundamental biophysical principle: proteins denature and aggregate when heated. Upon ligand binding, this thermal stability can shift, typically upward, offering an indirect but quantifiable measure of target engagement. Following ligand incubation, cells or lysates are exposed to a temperature gradient. Proteins that remain folded after heating stay in the soluble fraction, while denatured proteins aggregate and are removed by centrifugation. The soluble fraction is then quantified using Western blot, enzyme-linked immunosorbent assay (ELISA), or differential scanning fluorimetry (DSF). DSF employs environment-sensitive fluorescent dye that binds hydrophobic regions exposed during unfolding. The resulting fluorescence profile yields a melting temperature (T m ), which reflects the thermal stability of the protein. In most cases, ligand engagement increases the T m , offering a functional readout of stabilization. [7, 39] CETSA can be applied to intact cells, lysates, or even tissue extracts to assess target engagement in native-like environments while minimizing sample manipulation. [40] CETSA also supports an isothermal dose–response format, where thermal stabilization is measured at a single temperature with variable ligand concentration, generating dose-response curves which often correlate with half maximal inhibitory concentration (IC₅₀) values or K d values from orthogonal techniques. Alongside DSF, CETSA is widely used in drug discovery to quantify ligand–protein interactions across diverse affinity ranges. [11, 39, 40] One of CETSA’s key advantages is its ease of use. Unlike MS-based approaches, it does not require proteolytic digestion, complex instrumentation, or extensive data deconvolution. CETSA has proven to be adaptable to various MP classes, including SERCA2, PAR2, MCT4, and the adenosine A 2A receptor. [41-43] For instance, thermal stabilization of lauryl maltose neopentyl glycol (LMNG)-solubilized A 2A receptor upon ligand binding resulted in T m shifts of up to 9°C, while DDM-solubilized MCT4 exhibited a 10°C up-shift in response to incubation with its substrate. [42-43] Despite these promising results, CETSA’s utility for MPs remains constrained by key biochemical limitations. Success hinges on maintaining native protein conformation post-solubilization, which is highly detergent-sensitive. Moreover, the requirement for validated antibodies to detect thermally shifted proteins restricts assay scalability and limits the breadth of potential targets. These constraints explain the persistent underrepresentation of MPs in CETSA datasets. [7, 11, 41, 44] 3.2.2 Thermal Proteome Profiling Thermal Proteome Profiling (TPP) overcomes the throughput limitation of CETSA by integrating MS-based proteomics. [45, 46] Like CETSA, TPP applies a temperature gradient to intact cells or lysates treated with ligands, inducing protein unfolding ( Fig. 2B ). Soluble protein fractions are subsequently digested, optionally labeled with isobaric tandem mass tags (TMT), pooled, and analyzed via LC-MS/MS. This enables simultaneous generation of thermal shift curves for thousands of proteins, allowing broad identification of direct and indirect ligand targets. TPP also detects off-target interactions, aiding in the evaluation of drug specificity and safety. [47] Enhancements to the TPP workflow, such as TPP-compound concentration range and 2D-TPP, have improved sensitivity by profiling thermal shifts across drug concentration gradients and combined temperature-concentration conditions. [44] TPP has been widely applied to soluble proteomes across various cell types, establishing it as a cornerstone in proteomics and drug discovery. [7, 48] However, its reliance on protein abundance and MS ionization efficiency limits the detection of low-expression or poorly ionizing proteins, particularly many MPs. Despite these challenges, TPP has successfully profiled MPs such as CD45 and the Na⁺/K⁺-ATPase α-subunit in response to the cardiac glycoside ouabain. Comprehensive studies involving over 400 integral MPs across four human suspension cell lines revealed that proteins with higher numbers of transmembrane domains and α-helices generally exhibit greater thermal stability. This likely reflects stabilizing hydrophobic interactions within the lipid bilayer and highlights the importance of considering membrane topology when interpreting TPP-derived thermal profiles. [11, 13, 49] A notable advancement is cell-surface TPP (CS-TPP), introduced by Kalxdorf et al. (2021), enables selective thermal profiling of glycosylated extracellular MPs. The method utilizes hydrazide bead enrichment, whereby terminal sialic acid residues are oxidized to aldehydes, allowing for specific capture of surface-exposed glycoproteins. Using CS-TPP, the authors demonstrated ligand-induced stabilization of several MPs, including the chemokine receptor CCR5 and the monocarboxylate transporters MCT1 and MCT3. [13] This approach represents a major step forward in expanding TPP’s applicability to membrane-localized targets. Despite its strengths, TPP remains limited by its labor-intensive sample processing, which can involve up to 40 samples per experiment. [45, 50] To benchmark workflows, George et al. (2023) compared tandem mass tag-based data-dependent acquisition (TMT-DDA) and data-independent acquisition (DIA): while TMT-DDA identified more targets, DIA provided more consistent quantification. To further increase throughput, Ball et al. (2020) developed the isothermal shift assay (iTSA), which measures protein solubility at a single temperature using multiple replicates to improve statistical power. [50, 51] Although iTSA sacrifices full thermal curve resolution, it nearly doubled kinase target detection compared to standard TPP. More recently, single-tube TPP with uniform progression (STPP-UP) was developed by Zijlmans et al. (2023), introducing a single-tube temperature ramp up protocol that reduces sample handling. [52] Although less sensitive, STPP-UP offers a practical high-throughput option for early-stage drug screening. Together, these innovations and protocol optimizations underscore the growing potential of TPP not only for soluble proteomes but also for systematic, high-throughput exploration of MPs in pharmacological research. 3.3 Proteolytic Resistance Methods Similar to TPP, drug affinity responsive target stability (DARTS) was among the earliest methods to leverage ligand-induced stabilization, relying on protection from proteolysis and gel-based separation to identify protein targets. [53-54] While effective for soluble proteomes, DARTS is less suited for MPs due to the limited protease accessibility of transmembrane domains. [38, 55] Recent advances in MS have extended this concept through limited proteolysis mass spectrometry (LiP-MS), which maps ligand-induced proteolytic changes at the peptide level. [56, 57] LiP-MS workflows typically involve incubating protein lysates with small molecules, digesting with broad-spectrum proteases, and analyzing cleavage patterns via LC-MS/MS ( Fig. 2C ). Notably, Soste et al. (2023) applied LiP-MS to detect conformational changes in the GPCR ACKR3 upon ligand binding—indicating that even complex MPs can be interrogated without prior structural knowledge. [58] In a related development, proteome-wide elution profile learning for small-molecule activity (PELSA) mapped the binding site of the kinase inhibitor Lapatinib on ERBB2 by combining limited proteolysis with SEC, improving spatial resolution. [59] Most recently, thermostability-assisted LiP-MS (TALiP-MS) enhanced sensitivity by enriching thermally stabilized proteins prior to proteolysis, that is by heating drug treated cell lysates followed by centrifugation to remove aggregated heat-denatured proteins. This enrichment step increased target peptide detection by up to eightfold. [60] While TALiP-MS has yet to be applied to MPs, it represents a promising avenue for extending proteolytic-based profiling to challenging protein classes. 4. Shortcomings of Detergent in Membrane Proteins-Ligands Screenings Preserving MPs within their native lipid bilayers is a key objective for elucidating structure, function, and pharmacology, offering a more physiologically relevant foundation for drug discovery. Yet, this ideal is rarely achieved in high-throughput workflows, which continue to rely heavily on detergents to extract MPs for downstream analyses, particularly in MS-based target deconvolution. The disruptive impact of detergents on MP capture is well documented. For instance, Kawatkar et al. (2019) used the non-ionic detergent NP-40 in thermal stability assays to identify targets such as TSPO, SERCA2, and PAR2. However, they reported substantial variability: assay conditions required manual optimization for each target, and even then, thermal shift signatures were often weak or inconsistent, particularly for GPCRs and low-affinity ligands. [41] These inconsistencies arise not only from protein instability but also from the intrinsic behavior of detergents. Berlin et al. (2023) demonstrated that detergent micelles, particularly as they approach their cloud point—the temperature at which they begin to separate into distinct phases—can disrupt protein solubility and compromise the accuracy of thermal shift assays. This phase separation alters the micellar environment, potentially causing proteins to aggregate or misbehave in ways unrelated to thermal unfolding. [14] Rather than acting as passive solubilizers, detergents engage in complex, temperature-sensitive interactions with proteins, thereby distorting thermal profiles and elevating background signals. Recognizing these challenges, Ye et al. (2023) introduced a modified, lower-temperature TPP workflow. They demonstrated that both non-ionic (NP-40) and zwitterionic (CHAPS) detergents induce aggregation above 50°C, masking ligand-induced stabilization. By lowering thermal parameters, they recovered melting shifts and uncovered MP-ligand interactions that standard TPP had overlooked. [61] These findings highlight that optimization can mitigate, but cannot eliminate artifacts altogether. At the core of the problem is a fundamental truth: detergents are not inert. Their interactions with both proteins and ligands complicate interpretation, especially in assays that depend on subtle conformational changes. For thermal profiling platforms such as TPP and CETSA, which rely on precise stability signatures this presents a major challenge. As a result, detergent-based solubilization remains a bottleneck in reliably capturing MP–ligand interactions, underscoring the urgent need for alternative approaches that preserve native-like environments without compromising structural fidelity. 5. Implementing Membrane Mimetics in Membrane Protein-Ligand Screens Given the limitations of detergent, it is no surprise that membrane mimetics (MM) have emerged as promising alternatives, offering a means to maintain MPs in aqueous, native-like environments. These platforms, such as nanodiscs, Peptidiscs, and SMA lipid particles (SMALPs) replace the lipid bilayer, offering enhanced protein stability and functional fidelity. While historically leveraged in structural biology, MMs are now being adapted to functional assays, including MS-based ligand screening, where they show potential to overcome the destabilizing effects of detergents. [22, 23, 24] Accordingly, recent studies show these systems expanding the analytical reach of drug discovery by making MPs tractable to high-throughput, proteomics-based workflows. [10, 62-74] 5.1 Nanodiscs for Proteome-Level Analysis and Screening Nanodiscs are discoidal lipid bilayers stabilized by two amphipathic membrane scaffold proteins (MSPs), typically derived from apolipoprotein A1. Upon detergent removal with polystyrene beads, MSPs self-assemble with phospholipids and purified MPs into soluble complexes with tunable diameter size and lipid composition. This modularity makes nanodiscs particularly valuable for dissecting ligand–MP interactions under controlled, native-like conditions. [22, 63, 75] For example, nanodiscs have facilitated detailed analysis of substrate binding to cytochrome P450 enzymes, supporting precise kinetic measurements of drug metabolism and allosteric regulation. [75] Additionally, they have enabled mechanistic studies of the calcium-sensing receptor (CaSR), β₂-adrenergic receptor (β₂-AR), preserving ligand binding and G protein coupling. [62] Beyond functional assays, nanodiscs support a wide range of readouts, including scintillation proximity assays (SPA; Fig. 3A ) and fluorescence-based activation assays, highlighting the ability of this platform to preserve key conformational states critical for pharmacological interrogation. [63-65] However, the broader adoption of nanodiscs in high-throughput screening has been constrained by the need for target-specific reconstitution that often necessitates protein-specific optimization. To overcome this, proteome-wide solubilization strategies have emerged. Rather than reconstituting individual MPs, these approaches enable the assembly of entire membrane proteomes into soluble MP-libraries. This fundamental shift in MP reconstitution trades specificity for scalability, allowing functional interrogation at the systems level. [75] Early work by Marty et al. (2013) reported up to ~85% incorporation efficiency of E. coli MPs into nanodiscs. [66] Wilcox et al. (2015) extended the library approach to synaptosomal membranes, successfully preserving native receptor activity, as confirmed by functional activity assays. Furthermore, nanodisc libraries were integrated with MS to identify MP targets of Alzheimer’s-associated amyloid-β oligomers (AβOs). By UV-crosslinking AβOs to their MP targets, researchers enabled affinity purification followed by peptide-level identification via LC-MS/MS. The same platform was used to screen small-molecule inhibitors, where reduced crosslinking indicated competitive displacement. This ultimately led to identification of aurin tricarboxylic acid, a compound that reduced AβO binding by over 90% in both SMPL rat hippocampal neurons. [67] These advances have laid the foundation for scalable, MP-targeted screening in MM environments. [68, 69] Expanding on this, Mak et al. (2017) streamlined the workflow by assembling nanodiscs directly from whole-cell lysates, eliminating the need for membrane isolation. This innovation enabled the rapid generation of MP-enriched libraries from diverse sources, including HEK293 and red blood cells, broadening their applicability to both mechanistic studies and compound screening. [69] In parallel, Roy et al. (2015) used osteosarcoma cells, while making lipid adjustments, adding anionic lipids and cholesterol to enrich their library for GPCRs. [68] In sum, nanodiscs are steadily evolving from specialized structural tools into versatile, high-content screening platforms capable of bridging the gap between membrane proteomics and drug discovery. 5.2 Other Emerging Membrane Mimetic Platforms for Membrane Protein Screening Beyond nanodiscs, alternative MM systems are advancing the detergent-free solubilization and stabilization of MPs. Among these, the Peptidisc platform has emerged as a robust and scalable option. Unlike nanodiscs, which rely on MSP-assembled scaffolds of defined size, Peptidiscs form through the self-assembly of amphipathic peptides around MPs, enabling rapid reconstitution following detergent solubilization of the native membranes. This rapid peptide-based encapsulation avoids prolonged detergent-induced destabilization while accommodating a broad range of protein sizes and topologies. A notable advancement is the His-tagged Peptidisc, developed by Young et al. (2020), which enables downstream purification of the membrane proteome via nickel-affinity chromatography. [70] Initially applied to E. coli , it proved effective in stabilizing a broad spectrum of MPs. Subsequent studies extended the platform to more complex systems: Zhao et al. (2023) successfully reconstituted MPs from HeLa cells, while Antony et al. (2024) achieved proteome-wide stabilization in mouse liver tissue, collectively identifying over 800 unique MPs [71-73]. These results highlight the robustness of Peptidisc reconstitution across diverse species and tissue types, as well as its effectiveness in enriching MPs for downstream analysis. Building on this foundation, Jandu et al. (2024) introduced membrane mimetic–thermal proteome profiling (MM-TPP) workflow, integrating Peptidisc libraries into a TPP-based screening format ( Fig. 3B ). Using E. coli and mouse liver membrane proteomes, the study profiled thermal stabilization patterns induced by ATP and orthovanadate, enabling the detection of binding events to multiple ATP-binding MPs, including ABC transporters. Additionally, the analysis revealed off-target thermal effects across the membrane proteome related to ADP- and AMP-binding MPs, likely arising from ATP metabolism. [10] While the promiscuity of ATP limits specificity, the results validate Peptidisc as an effective interface between MPs and proteome-scale screening platforms, establishing a proof-of-principle for future MS-based ligand discovery in MM environments. A third, increasingly adopted MM system is the SMA copolymer, which extracts MPs by solubilizing intact nanoscale membrane fragments termed SMALPs. These particles preserve native lipid–protein interactions and eliminate the need for detergents entirely. [63, 76, 77] While mostly applied for structural studies, SMALPs are now being adapted for proteomics and ligand screening workflows. For example, Sharma et al. (2021) showed that SMALP-reconstituted tetraspanins such as CD81 and CD53 retained native epitope accessibility and ligand-binding kinetics comparable to those in intact membranes, as confirmed by surface plasmon resonance. [74, 78] These results suggest that SMALPs preserve essential functional properties of MPs, making them a promising platform for downstream ligand characterization. However, challenges remain. Kamilar et al. (2023) reported that SMALPs can exhibit heterogeneous morphologies, forming assemblies ranging from single particles to mixed micelles, features that may complicate interpretation in high-throughput workflows. [79] Despite this, the utility of SMALPs in proteome-scale experimentation continues to grow. Mueller et al. (2023) benchmarked several SMA variants, including SMA 200, 300, and 502-E, against conventional detergents such as DDM and LMNG for solubilizing membrane proteomes from HEK293 cells. Across a broad range of MP classes, SMA-based systems matched or exceeded the extraction efficiency of detergents. [80] Expanding on this, Brown et al. (2025) used a suite of formulations to generate libraries containing over 2,000 mammalian MPs, fully compatible with MS-based workflows ( Fig. 3C ). [27] Together, these emerging platforms offer promising avenues for bridging the gap between MPs and MS-based ligand discovery. Peptidiscs enable affinity enrichment and scalability, while SMALPs preserve native lipid–protein interactions. Importantly, both are expanding the toolkit for capturing ligand–protein interactions. Collectively, these technologies are helping to make MPs—long considered challenging targets—more tractable for high-throughput screening and functional analysis. 6. Conclusion and Future Perspectives Despite their central role in cellular function, MPs have historically been underrepresented in ligand screening campaigns, largely due to their low abundance, structural complexity, and incompatibility with high-throughput workflows. However, these limitations are increasingly being addressed through technological innovations that preserve native MP structure and function at scale. MS, long established for soluble protein analysis, is now rapidly evolving into a platform for MP-ligand discovery. When paired with MM systems, MS can interrogate dynamic protein-ligand interactions within environments that more closely resemble native membranes, overcoming previous barriers related to solubility and low abundance. This convergence of MM platforms with affinity- and stability-based MS assays holds promise for improving MP-ligand screening methods. Rather than being excluded, MPs can now be systematically integrated into high-throughput workflows. Importantly, pairing direct binding approaches such as AS-MS with activity-based assays can enable drug discovery pipelines that not only identify ligand binding events, but also establish mechanistic links to functional outcomes. [81] This integration enhances confidence in target relevance and therapeutic potential, particularly valuable when screening MPs with poorly understood biology. However, it is equally important to recognize the caveats of these MS-based assays: several methods (e.g., CETSA, TPP, MSBA) are known to suffer from off-target effects, nonspecific interactions, or background signal interference, issues that are magnified in membrane environments. Additionally, ligand degradation or instability may also confound data interpretation. These challenges highlight the value of hybrid strategies and orthogonal validation. For instance, combining MSBA with AS-MS, or integrating TPP with Peptidisc-based solubilization, can strengthen target validation and improve mapping of binding sites. Multi-method workflows exploiting different assay mechanisms reduce the risk of false positives. Looking forward, the next generation of technologies will likely further accelerate MP-targeted ligand discovery. Cross-linking mass spectrometry (XL-MS) can define spatial constraints of binding sites. AI-driven deconvolution of complex MS datasets promises to streamline hit identification and target annotation. Integration with cryo-EM may enable structurally guided ligand design directly from MS-enriched samples, while single-cell proteomics offers a route to interrogate low-abundance MPs across heterogeneous tissues or disease states. Together, these innovations will not only expand the scope of MP drug discovery but may ultimately redefine what is considered “screenable” in the human proteome. Acknowledgements This work was supported by the Canadian Institutes of Health Research Project Grant 20R34019. Competing Interests: The authors declare no competing interests associated with the manuscript. Figure Legend Figure 1. Schematic Overview of Affinity-Based Mass Spectrometry Approaches for Ligand–Protein Interaction Discovery (A) Schematic of Affinity-Selection Mass Spectrometry (AS-MS). A small molecule library is incubated with a tagged target protein immobilized on a filter or a matrix (e.g., Ni-NTA for His-tagged proteins). Unbound and weakly bound compounds are removed via wash steps, vacuum filtration, ultracentrifugation, or size exclusion chromatography (SEC). Bound ligands are then eluted using a denaturing solvent such as acetonitrile, methanol, or urea. The ligand mixture is further separated by SEC, and individual fractions are analyzed by LC/MS to identify protein-binding compounds. (B) Schematic of Mass Spectrometry Binding Assays (MSBA). This approach uses three parallel incubations to distinguish specific from non-specific binders. First, a known unlabeled reporter ligand is incubated with the protein-matrix to confirm target engagement via LC/MS (control). Second, the test compound library is incubated with the protein to capture all potential binders (total binding). Third, the compound library is incubated with the protein in the presence of excess reporter ligand to competitively block specific binding sites, revealing non-specific binders (non-specific binding). Subtraction of the non-specific binding signal from the total binding yields the specific binders within the compound library. Key reagents and steps are indicated in the schematic, including ligand addition, elution, and MS readouts. Figure 2. Schematic Overview of Stability-Based Mass Spectrometry Approaches for Ligand–Protein Interaction Discovery (A) Schematic of Stability of Proteins from Rates of Oxidation (SPROX). The protein of interest is incubated with hydrogen peroxide and a denaturant such as urea, in both control and ligand-treated conditions. In the absence of ligand, protein unfolding exposes buried methionine residues, which are subsequently oxidized. After digestion with trypsin, oxidized peptides are analyzed by LC/MS, with oxidation-induced mass shifts reflecting the protein’s denaturation profile. In the presence of ligand, binding can stabilize the protein of interest, altering its unfolding trajectory and resulting in a distinct methionine oxidation pattern. Comparing these profiles allows identification of ligand-induced stabilization. (B) Schematic of Thermal Proteome Profiling (TPP). The proteome of interest is heated at different temperatures using a thermal cycler, causing thermally unstable proteins to eventually denature. The sample is then centrifuged at high speed, separating denatured proteins, which pellet, from soluble, non-denatured proteins, which remain in the supernatant. The supernatant is analyzed by LC/MS, only detecting the non-denatured proteins. In the ligand-treated condition, ligand binding may thermally stabilize specific proteins, preventing their denaturation and allowing them to remain in the soluble fraction. These stabilized proteins are detected by LC/MS, and the difference in protein abundance between conditions enables the inference of ligand–protein interactions through thermal stabilization. (C) Schematic of Limited Proteolysis Mass Spectrometry (LiP-MS). The protein of interest is treated with a non-selective protease in both control and ligand conditions. In the absence of ligand, accessible regions are digested, generating characteristic peptide fragments detected by LC/MS. Ligand binding can occlude specific protease cleavage sites, leading to altered digestion patterns. Comparative analysis of these peptide profiles reveals ligand-induced conformational protection and identifies potential binding regions. Figure 3. Membrane Mimetic Strategies for Membrane Protein Ligand Interaction Studies and for Membrane Proteome Profiling (A) Schematic of Nanodisc Scintillation Proximity Assay (SPA). A His-tagged membrane protein (MP) of interest is solubilized in detergent and reconstituted into nanodiscs following detergent removal, facilitated by adsorbent beads. The nanodisc-reconstituted MP is immobilized on scintillation-coated beads via a His-tag. A radiolabeled ligand is then incubated with the immobilized nanodisc-MP. Upon binding to the MP, proximity to the scintillant surface enables detection of ligand–protein interactions via emitted light. This setup allows for direct, label-free quantification of binding events in a membrane-like environment. (B) Schematic of Membrane Mimetic Thermal Proteome Profiling (MM-TPP). MPs are solubilized in detergent and reconstituted with Peptidisc using a centrifugal unit to remove detergent and facilitate self-assembly. The reconstituted membrane proteome is incubated with or without a ligand (ATP), then subjected to heat-induced denaturation using a thermal cycler. Following ultracentrifugation, thermally unstable proteins pellet, while stabilized proteins remain in the supernatant. Peptides from the pellet or supernatant are digested and analyzed by LC/MS. Ligand-induced thermal stabilization is inferred from shifts in the abundance of specific MPs between control and ligand-treated conditions. (C) Schematic of styrene maleic acid lipid particle (SMALP) membrane proteome profiling. MPs are directly extracted from crude membrane preparations using styrene–maleic acid (SMA) copolymer, which solubilizes lipid bilayers without the use of detergent, preserving native protein–lipid interactions. To separate proteins from the SMA copolymer and associated lipids, the solubilized proteome is subjected to precipitation using a 3:1 ratio of methyl tert-butyl ether and methanol. 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Collection PROTEOMICS Keywords biochemistry drug discovery lipid bilayer membrane mimetics membrane proteins Authors Affiliations Jonathon Lambos UBC View all articles by this author Ashim Bhattacharya UBC View all articles by this author Mohammed Al-Seragi UBC View all articles by this author Franck Duong 0000-0001-7328-6124 [email protected] UBC View all articles by this author Metrics & Citations Metrics Article Usage 685 views 369 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Jonathon Lambos, Ashim Bhattacharya, Mohammed Al-Seragi, et al. Expanding the Reach of Membrane Protein-Ligand Interaction Studies through the integration of Mass Spectrometry and Membrane Mimetics. Authorea . 13 August 2025. DOI: https://doi.org/10.22541/au.175508838.89786308/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. 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