A High-Throughput Fluorometric Assay for Lipid–Protein Binding

CHAPTER ONE

A High-Throughput Fluorometric Assay for Lipid–Protein Binding W. Cho*,†,1, Y. Hu*, K. Baek†, H. Kim* *University of Illinois at Chicago, Chicago, IL, United States † Kyung Hee University, Yongin, South Korea 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Fluorescence Quenching Assay 2.1 Assay Design and Optimization 2.2 Preparation of Quencher Lipid Vesicles 2.3 Preparation of FP-Tagged Proteins 2.4 Plate Reader-Based Quantitative Lipid Specificity Assay 2.5 Plate Reader-Based Membrane-Binding Affinity Assay 2.6 Spectrofluorometric Membrane-Binding Affinity Assay 2.7 High-Throughput Screening of Membrane-Binding Inhibitors 3. Data Analysis 3.1 Specificity Data Analysis 3.2 Binding Data Analysis 3.3 Inhibition Data Analysis Acknowledgments References

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Abstract An increasing number of intracellular and extracellular proteins are shown to interact with membrane lipids under physiological conditions. For rapid and robust quantitative measurement of lipid–protein interaction, we developed a sensitive fluorescence quenching-based assay that is universally applicable to all proteins and lipids. The assay employs fluorescence protein (FP)-tagged proteins whose fluorescence emission intensity is decreased when they bind vesicles containing quenching lipids. This simple assay can be performed with a fluorescence plate reader or a spectrofluorometer and optimized for different proteins with various combinations of FPs and quenching lipids. The assay allows a rapid, sensitive, and accurate determination of lipid specificity and affinity for various lipid-binding proteins, and high-throughput screening of molecules that modulate their membrane binding.

Methods in Enzymology, Volume 583 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.09.004

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2017 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Membrane lipids regulate and coordinate a wide variety of biological processes by interacting with diverse proteins (Cho, 2006; van Meer, Voelker, & Feigenson, 2008). It has been well documented that the lipid–protein interaction is important for the structure and function of transmembrane proteins that constitute about one-third of human proteome (Barrera, Zhou, & Robinson, 2013; Lee, 2011; van Meer et al., 2008). In addition, many intracellular and extracellular proteins that are involved in various processes, including cell signaling, membrane trafficking, blood coagulation, and viral infection, are recruited to cell membranes through lipid–protein interactions (Cho, 2006; Cho & Stahelin, 2005; DiNitto, Cronin, & Lambright, 2003; Lemmon, 2008). Most of these proteins contain lipid-binding domains or motifs that mediate membrane binding either through specific recognition of lipid headgroups or by nonspecific electrostatic and hydrophobic interactions with membrane lipids (Cho & Stahelin, 2005; DiNitto et al., 2003; Lemmon, 2008). Recent bioinformatics calculations have predicted that a large portion of cellular proteins may be able to interact with membrane lipids under physiological conditions because they have structural features necessary for membrane binding (Bhardwaj, Stahelin, Zhao, Cho, & Lu, 2007; Silkov et al., 2011). Consistent with the prediction, recent genome-wide studies of protein domains, including PSD95, Dlg1, ZO-1 (PDZ) and Src-homology 2 (SH2) domains, have demonstrated that a large number of these domains bind membrane lipids with high affinity and specificity (Bhardwaj, Stahelin, Langlois, Cho, & Lu, 2006; Chen et al., 2012; Feng & Zhang, 2009; Park et al., 2016; Sheng et al., 2012, 2014; Silkov et al., 2011; Zimmermann, 2006). These results suggest that the number of membrane-binding intracellular and extracellular proteins is much larger than originally suspected and that membrane binding may be an important regulatory step for these proteins. A rapid, sensitive, and quantitative assay for membrane–protein interaction is thus necessary for further identification and characterization of rapidly growing families of membrane-binding proteins and for identification of small molecules modulating their membrane interaction. Membrane binding of soluble proteins has been measured by various biochemical and biophysical methods (Cho, Bittova, & Stahelin, 2001; Narayan & Lemmon, 2006). Sedimentation assays using lipid vesicles (Rebecchi, Peterson, & McLaughlin, 1992) or lipid-coated beads (Kim,

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Lichtenbergova, Snitko, & Cho, 1997) have been most commonly used to assess membrane binding of proteins. However, difficulties associated with accurately quantifying membrane-bound vs free proteins and variable pelleting efficiency associated with different lipid vesicles have limited their utility. The lipid overlay assay has been commonly used due to its convenience but it suffers from many drawbacks, including low sensitivity, poor reliability, and an inability to yield quantitative information (Dowler, Kular, & Alessi, 2002). Also, lipids are presented in a poorly defined, nonphysiological state in this assay. The surface plasmon resonance (SPR) analysis allows robust quantitative analysis of membrane–protein interactions and has thus been a mainstay in biophysical characterization of membrane-binding proteins (Stahelin, 2013; Stahelin & Cho, 2001). Although this assay offers many advantages, including high sensitivity, no requirement for protein labeling, and an ability to provide kinetic information, it also has drawbacks, including the necessity of expensive instrumentation and rigorous controls to eliminate nonspecific binding, uncertainty about the physical nature of lipids coated on the sensor chip, and binding measurements under nonequilibrium conditions. Due to their high sensitivity, various fluorescence techniques have been extensively employed to monitor membrane–protein interaction. Most commonly, an increase (Kraft, Garrido, Leiva-Vega, & Romero, 2009), quenching (Dua, Wu, & Cho, 1995), or fluorescence resonance energy transfer (Bazzi & Nelsestuen, 1987; Nalefski, Slazas, & Falke, 1997) of Trp fluorescence is monitored during membrane– protein binding. Although rapid and convenient, these methods are not generally applicable to all proteins because many proteins do not have Trp on their membrane-binding surfaces. Although Trp can be genetically introduced to the membrane-binding surface to a protein, this can dramatically change its membrane-binding property (Sumandea, Das, Sumandea, & Cho, 1999). Alternatively, the protein can be labeled with an organic fluorophore, but this approach is limited by experimental inconvenience and the relative low yield of chemical modification (Liu et al., 2014; Yoon, Lee, Kurilova, & Cho, 2011). Fluorescence anisotropy (Miao et al., 2010) and fluorescence correlation spectroscopy analyses (Rusu, Gambhir, McLaughlin, & Radler, 2004) have been also used to measure membrane–protein interaction but with limited applications. Most important, none of these methods are suitable for robust and universal high-throughput analysis. To overcome these technical limitations and obstacles, we developed a high-throughput membrane-binding assay that is based on

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fluorescence quenching of fluorescence proteins (FPs), such as enhanced green FP (EGFP), fused to a membrane-binding protein by a dark quencher-containing lipid, such as N-dimethylaminoazobenzenesulfony l-phosphatidylethanolamine (dabsyl-PE), incorporated in lipid vesicles (Cho, Kim, & Hu, 2016; Kim, Afsari, & Cho, 2013). Since FPs do not have affinity for membrane lipids (Kim et al., 2013), an FP tag does not affect the membrane binding of a diverse group of proteins tested so far. Furthermore, FPs offer an additional advantage of stabilizing the fused protein or domain (Park et al., 2016). Through the custom selection of an FP and a dark quencher from widely available collections, this simple and rapid assay can be optimized for sensitive, accurate, and reproducible quantitative determination of lipid affinity and specificity of diverse proteins as well as for high-throughput screening of small molecules that can modulate their membrane binding.

2. FLUORESCENCE QUENCHING ASSAY 2.1 Assay Design and Optimization As described earlier, intrinsic Trp fluorescence cannot be utilized for a sensitive, universal membrane-binding assay. Also, chemical labeling of proteins, although offering higher sensitivity, suffers from low labeling yield and experimental inconvenience. Our strategy for the universal fluorescence assay is to genetically render favorable fluorescence properties to all proteins by expressing them as FP-fusion proteins and then quantify their membrane binding by monitoring the decrease in FP fluorescence as the proteins bind the lipid vesicles incorporating a nonfluorescent quenching dye (dark quencher; see Fig. 1A). FP-fusion proteins have been extensively used to monitor the subcellular localization, dynamics, and interactions of numerous cellular proteins (Miyawaki, 2011; Shaner, Steinbach, & Tsien, 2005) and many FP tags are known to have a minimal effect on the membrane-binding properties of fusion partner proteins when attached either their N- or C-terminus with an appropriate linker (Kim et al., 2013; Park et al., 2016). As is the case with other fusion tag proteins, FP tags also stabilize the partner proteins, especially when they are expressed as isolated protein domains or truncated proteins, thereby allowing higher expression yields of recombinant proteins (Park et al., 2016). EGFP is a preferred FP because it is the most commonly used FP with relatively high quantum yield and with a lesser effect on protein aggregation than other FPs (Miyawaki, 2011;

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Fig. 1 Principles and components of our fluorescence quenching-based membranebinding assay. (A) The general strategy of the assay. To the solution of an FP-fusion protein (e.g., Akt1-PH-EGFP) is added vesicles containing their favorite lipids (e.g., PtdIns (3,4,5)P3) and a dark quencher, such as dabsyl-PE (shown in black). As the protein approaches the membrane surface, the fluorescence emission intensity of FP is greatly reduced due to quenching by dabsyl-PE, allowing quantitative analysis of membrane– protein interaction. (B) The chemical structure of dabsyl-PE. The quencher moiety is highlighted by the dotted oval. (C) The chemical structures of dabsyl-PE derivatives with extended linkers. Typically, R1 ¼ palmitoyl, R2 ¼ oleoyl, n ¼ 2–5. (D) The absorption spectra of dabsyl-PE, BHQ1-PE, and QSY7-PE (all 60 nM) in chloroform. (E) Fluorescence excitation (Ex) and emission (Em) spectra of EGFP.

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Shaner et al., 2005). In the case higher sensitivity is required for the assay, a brighter FP, such as mNeonGreen (Shaner et al., 2013), can be substituted for EGFP. Dabsyl-PE (Fig. 1B) has been primarily used as a quenching lipid for routine assays because it is easy and inexpensive to prepare in large quantities. Because its absorption spectrum (Fig. 1D) overlaps more with the excitation spectrum than with the emission spectrum of EGFP (Fig. 1E), however, it is not ideal for the assay employing EGFP-fusion proteins. For high quenching efficiency and assay sensitivity, one can employ black hole quencher® 1 (BHQ1)-PE or QSY®7-PE for dabsyl-PE as a quenching lipid (Fig. 1D). BHQ1-PE typically allows >30% improvement over dabsyl-PE in EGFP quenching efficiency. QSY7-PE (absorption maximum at  570 nm; see Fig. 1D) is better suited for YFP (λEx ¼ 514 nm; λEm ¼ 527 nm) and YFP quenching can be measured with λEx ¼ 510 nm and λEm ¼ 540 nm. Despite their improved spectral properties, however, general use of BHQ1-PE and QSY7-PE is limited because they are expensive to prepare in large quantities. Another factor in assay design is the quenching efficiency that depends on the distance between the lipid quencher and the FP. Ideally, an FP tag should be located close enough to the membrane-binding surface of the host protein to yield >40% of quenching upon membrane binding. One can achieve higher quenching yield by changing the location of the FP tag (i.e., N- or C-terminal) or the length of the linker between the FP and the host protein. Another way to improve the quenching efficiency is to extend the hydrophilic linker between PE and the quencher moiety of the quenching lipid (e.g., see Fig. 1C) so that the quencher can stretch further into the solution.

2.2 Preparation of Quencher Lipid Vesicles The lipid composition of vesicles and the mole% of a quenching lipid can be adjusted according to the purpose of the assay. All PE-based quenching lipids are anionic lipids and might nonspecifically increase the vesicle binding of proteins with cationic membrane-binding surfaces. For accurate determination of membrane-binding affinity (or specificity) of a protein under physiological conditions, the concentration of a quenching lipid must be thus kept as low as possible (i.e., just high enough to allow robust quenching). Typically, 40% of FP quenching is sufficient for robust and reproducible binding analysis. For initial assessment of membrane-binding

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affinity of a large number of proteins and high-throughput screening of small molecule, one can increase the concentration of the quenching lipid to maximize the FP quenching efficiency. In all cases, background correction with lipid vesicles containing primarily the quenching lipid is recommended. For example, for phosphatidylinositol-3,4,5-trisphosphate (PtdIns(3,4,5)P3)-binding proteins that are assayed with 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC)/dabsyl-PE/PtdIns (3,4,5)P3 (92:5:3) vesicles, background correction with POPC/dabsyl-PE (95:5) is appropriate. Here, POPC is used as a bulk lipid because most proteins do not bind PC. Large unilamellar vesicles (LUVs) are more uniform and stable than small unilamellar vesicles (SUVs) and they are thus more suitable for accurate binding measurements. The uniformity of vesicles is also important for quenching efficiency because the presence of vesicles lacking quenching lipids would lower the overall quenching efficiency by sequestering FP-tagged proteins. However, it takes much longer to prepare LUVs through extrusion, which becomes a major limiting factor for high-throughput screening that requires a large amount of vesicles. For high-throughput screening, SUVs are thus preferred to LUVs. Due to their low stability, SUVs should be used within a few hours of preparation. 1. POPC (Cat. No. 850457), 1-palmitoyl-2-oleoyl-sn-glycero-3phosphoethanolamine (POPE; Cat. No. 850757), 1-palmitoyl-2oleoyl-sn-glycero-3-phosphoserine (POPS; Cat. No. 840034), or soy phosphatidylinositol (PtdIns, Cat. No. 840044) (all from Avanti Polar Lipids) are dissolved in the highest grade oxygen-free chloroform to yield 10 mg/mL stock solutions and stored in Teflon-sealed vials at 20°C. All organic solvents including chloroform (Cat. No. C607-4), methanol (Cat. No. 53816), triethylamine (Cat. No. O4885-1), and dichloromethane (Cat. No. D37-1) are from ThermoFisher (Waltham, MA, USA). 2. 1,2-Dipalmitoyl derivatives of phosphoinositides (PtdInsPs), including PtdIns(3,4,5)P3 (Cayman Chemical, Cat. No. 64920), are dissolved in the highest grade oxygen-free chloroform/methanol/water (3:3:1 v/v/v) to yield 0.5 mg/mL solutions and also stored in Teflon-sealed vials at 20°C. 3. Dabsyl-PE is synthesized from POPE as follows: POPE (50 mg) is dissolved in chloroform (2 mL) and the solution is added to a solution of dabsyl chloride (Tokyo Chemical Industry, Cat. No. D1382; 22.6 mg) and triethylamine (0.2 mL) in chloroform (5 mL). The mixture is stirred for 6 h at room temperature in the dark and the solvent

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removed in vacuo. The residue is dissolved in dichloromethane/ methanol (9:1) and purified by silica column chromatography using the same solvent mixture as eluent. The solvent is evaporated in vacuo to afford dabsyl-PE as an orange solid. A stock solution of dabsyl-PE in chloroform (10 mg/mL) is prepared and stored in a Teflon-sealed vial at 20°C. 4. BHQ1-PE and QSY7-PE are synthesized as follows: 8 mg of POPE and 5 mg of QSY7 carboxylic acid succinimidyl ester (ThermoFisher, Waltham, MA, USA, Cat. No. Q10193) or BHQ1 carboxylic acid succinimidyl ester (LGC Biosearch Technologies, Cat. No. BHQ1000S-5) are mixed in 800 μL of chloroform and 200 μL of triethylamine. The reaction and purification are performed as described for dabsyl-PE. 10 mg/mL stock solutions of BHQ1-PE and QSY7-PE in chloroform are prepared and stored in Teflon-sealed vials at 20°C. 5. LUVs (100-nm diameter) are prepared by mixing the lipid solutions in chloroform according to the final lipid composition (e.g., POPC/ dabsyl-PE/PtdIns(3,4,5)P3 ¼ 92:5:3) and the solvent is evaporated under the gentle stream of nitrogen gas. 20 mM Tris (ThermoFisher, Waltham, MA, USA, Cat. No. 142521) buffer, pH 7.4, containing 0.16 M NaCl (ThermoFisher, Waltham, MA, USA, Cat. No. 156697) is added to the lipid film to adjust the final lipid concentration, and the mixture is vortexed for 1 min, and sonicated in a sonicating bath for 1 min to break multilamellar vesicles. LUVs are prepared by multiple extrusion through a 100-nm polycarbonate filter (Avanti, Cat. No. 610005) using a MiniExtruder (Avanti, Cat. No. 610000). 6. SUVs (<15-nm diameter) are prepared by mixing the lipid solutions in chloroform according to the final lipid composition and the solvent is evaporated under the gentle stream of nitrogen gas. 20 mM Tris buffer, pH 7.4, containing 0.16 M NaCl is added to the lipid film to adjust the final lipid concentration, and the mixture is vortexed for 1 min, and sonicated using a Branson Sonifier until the solution becomes clear.

2.3 Preparation of FP-Tagged Proteins For most proteins, N-terminal and C-terminal FP tags have essentially the same effect: i.e., they improve the protein expression yield without affecting membrane-binding properties of the fusion partner proteins (Park et al., 2016). Depending on the structure of the protein, however, N-terminal and C-terminal FP tags may differentially affect the structure and function of the proteins and also the quenching efficiency of FP in the assay. The

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location of the FP tag and the sequence and length of the linker between FP and the host protein should thus be adjusted to optimize the assay condition. FP-tagged proteins are typically expressed with a His6 tag and purified by affinity chromatography using a Ni-chelate column. For some proteins, most notably electrostatically neutral proteins, the His6 tag may significantly enhance the binding of a neutral to anionic protein to anionic vesicles. In such a case, the His6 tag should be enzymatically removed after affinity purification or the assay should be performed at slightly elevated pH (e.g., pH 7.8) to ensure that all His side chains are deprotonated. Also, the protein can be expressed as a glutathione-S-transferase (GST)-tagged protein, purified by glutathione-based affinity chromatography, and the GST tag removed before the assay. A typical procedure for bacterial expression of EGFP- and His6-tagged protein is as follows: 1. 500 mL of Luria (BD, Cat. No. 244610) broth containing 50 μg/mL kanamycin (ThermoFisher, Waltham, MA, USA, Cat. No. 155479) or 100 μg/ mL ampicillin (ThermoFisher, Waltham, MA, USA, Cat. No. 160561) is inoculated with BL21 RIL colonies expressing each FP-tagged protein. 2. Cells are allowed to grow in the medium at 37°C until an absorbance at 600 nm reaches 0.6. 3. Protein expression is induced with the addition of 100 μM isopropyl 1-thio-β-D-galactopyranoside (Gold Biotechnology, St. Louis, MO, USA, Cat. No. 12481C5), and cells are moved to a 25°C shaker for 14 h incubation. 4. Cells are harvested by centrifugation (2500  g for 10 min at 4°C), and the pellet is resuspended in 20 mL of the lysis buffer (50 mM Tris– HCl, 300 mM NaCl, 10 mM imidazole (ACROS Organics, Cat. No. B0141064), and 10% (v/v) glycerol (ThermoFisher, Waltham, MA, USA, Cat. No. 155081), pH 7.9). 5. The solution is sonicated for 5 min with 30-s intervals and the mixture is centrifuged for 30 min (39,000  g at 4°C). The supernatant is transferred to a 50-mL Falcon tube and 1 mL of Ni-NTA agarose (Qiagen, Cat. No. 151032765) is added to it. 6. The supernatant is allowed to equilibrate with the resin for 30 min at 4°C with gentle mixing. The supernatant is poured onto a column, and the resin is washed with 50 mL of 50 mM Tris buffer, pH 7.4, containing 20 mM imidazole and another 50 mL of 50 mM Tris buffer, pH 7.4, containing 40 mM imidazole. The protein is eluted using 50 mM Tris buffer, pH 7.4, with 300 mM imidazole. 7. The purity of the eluted protein is checked by sodium-dodecylsulfate (ThermoFisher, Waltham, MA, USA, Cat. No. BP166-500) gel

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electrophoresis using an 18% polyacrylamide gel, and the protein concentration is determined during the Bradford reagents (BioRad, Hercules, CA, USA, Cat. No. 5000006). The protein is frozen in liquid nitrogen and stored at 20°C.

2.4 Plate Reader-Based Quantitative Lipid Specificity Assay Many membrane-binding proteins coincidently interact with both bulk lipids and a specific lipid under physiological conditions (Cho & Stahelin, 2005). For example, most of pleckstrin homology (PH) domains interact with multiple anionic lipid molecules, most notably phosphatidylserine (PS), via their membrane-binding surface and a PtdInsP molecule in their lipid-binding pocket (Lucas & Cho, 2011; Manna, Albanese, Park, & Cho, 2007; Manna et al., 2008; Stahelin et al., 2003). For accurate, physiologically relevant determination of lipid specificity, a two-step assay is thus recommended. First, any requirement for a bulk lipid for a protein is determined using POPC/dabsyl-PE/L1 (95x:5:x) (L1 ¼ PS, cholesterol, etc.; x ¼ 0–30 mole%) vesicles. Then, using POPC/dabsyl-PE/L1/L2 (75 x:5:20:x) (L2 ¼ PtdInsP, phosphatidic acid, diacylglycerol, etc.; x ¼ 0–10 mole%; 20 mole% is an arbitrary concentration for a bulk lipid) vesicles, specificity (headgroup or acyl chain) of its lipid-binding pocket can be determined. This two-step assay is preferred to a single-step assay employing, e.g., POPC/dabsyl-PE/L (70:5:25) (L ¼ all lipids) because it is not physiologically meaningful to directly compare the affinity for bulk lipids, such as PS, and that for signaling lipids, such as PtdIns(3,4,5)P3, which exist in much lower concentrations in cell membranes. 1. A fixed concentration of protein (e.g., 100 nM) is added to each well of a Corning nontreated black polystyrene 96-well plate (ThermoFisher, Waltham, MA, USA, Cat. No. 07-200-590). 2. To each row vesicles with varying lipid composition (e.g., POPC/ dabsyl-PE/L1 (95x:5:x) (L1 ¼ any signaling lipid such as PtdInsP, phosphatidic acid, and etc.; x ¼ 0–10 mole%)) are added. The total lipid concentration (e.g., 10 μM) is kept constant and the final volume of the mixture in the assay buffer is adjusted to 200 μL. 3. Other rows are filled with different lipid mixtures (e.g., POPC/dabsylPE/L2 (95 x:5:x), POPC/dabsyl-PE/L3 (95 x:5:x), etc.). 4. One row is reserved for background correction for nonspecific quenching using the same protein and POPC/dabsyl-PE (95:5) vesicles. 5. The plate is incubated at 25°C with extremely gentle shaking for 5 min.

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6. The decrease in EGFP fluorescence emission intensity at 509 nm is monitored with excitation set at 488 nm using a fluorescence plate reader. Excitation and emission wavelengths are adjusted for other FPs.

2.5 Plate Reader-Based Membrane-Binding Affinity Assay Once lipid specificity of a protein is known, its affinity for the favorite lipid vesicles, such as POPC/POPS/PtdIns(3,4,5)P3/dabsyl-PE (72:20:3:5), is determined by varying the total concentration of the same lipid vesicles with a fixed concentration of the protein. An alternative way is to employ the fixed total concentration of lipid vesicles and varying concentrations of the protein. This approach is useful when the nonspecific quenching becomes significant because of low quenching efficiency of specific binding. 1. To each well of a given row of a 96-well plate a fixed concentration of protein (e.g., 100 nM) and an increasing concentration of lipid vesicles with fixed composition (e.g., 0–150 μM of POPC/PtdIns(3,4,5) P3/dabsyl-PE (92:3:5) vesicles) are added. The final volume of the assay mixture is adjusted to 200 μL. 2. Multiple (i.e., triplicate) rows are filled with the same protein and lipid mixtures for multiplex determination. 3. One row is selected for background correction for nonspecific quenching using the same protein and dabsyl-PE vesicles (e.g., POPC/dabsyl-PE (95:5) vesicles). 4. The plate is incubated at 25°C with extremely gentle shaking for 5 min. 5. The decrease in EGFP fluorescence emission intensity at 509 nm is monitored with excitation set at 488 nm using a fluorescence plate reader. Excitation and emission wavelengths are adjusted for other FPs.

2.6 Spectrofluorometric Membrane-Binding Affinity Assay The cuvette-based spectrofluorometric assay is recommended when accurate and robust quantification of membrane affinity of proteins is required for biophysical analysis (e.g., binding energy calculation). 1. 10–20 μL of the protein stock solution is added to 2 mL of the assay buffer equilibrated at 25°C to the final concentration (typically 100 nM) depending on the Kd value for the protein and the particular vesicles (e.g., POPC/PtdIns(3,4,5)P3/dabsyl-PE (92:3:5) vesicles). The protein solution is transferred to a 3-mL (or smaller) quartz cuvette in a thermostated (25°C) spectrofluorometer.

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2. The equilibrated value of EGFP fluorescence emission intensity at 509 nm is monitored with excitation set at 488 nm after 5 μL incremental addition of the lipid vesicle solution. 3. Lipid addition is continued until binding reaches saturation. 4. The same experiment is repeated with dabsyl-PE vesicles (e.g., POPC/ dabsyl-PE (95:5) vesicles) for background correction for nonspecific quenching.

2.7 High-Throughput Screening of Membrane-Binding Inhibitors This assay has been also used to screen small molecules that specifically modulate the membrane binding of various proteins through competition analysis (Cho et al., 2016; Kim et al., 2013). In most cases, molecules are screened for inhibitory activity by monitoring dequenching of FP fluorescence as a consequence of membrane unbinding of proteins. Since the inhibition assay monitors the increase of FP fluorescence, it allows high sensitivity. It should be noted that some active small molecules are fluorescent and may have spectral overlap with FP and dabsyl-PE. Therefore, rigorous controls and background correction are necessary to minimize false-positive hits. The assay is typically run in two steps: an initial high-throughput assay using a single-point lipid determination (e.g., a fixed vesicle concentration giving a half-maximal quenching of EGFP in the absence of an inhibitor), followed by a more accurate determination with multiple lipid concentrations. The assay can also be used to screen molecules that allosterically enhance the membrane binding of a protein. In this case, increased quenching of FP signals (i.e., a further decease in fluorescence intensity) is monitored. 2.7.1 High-Throughput Inhibitor Screening Protocol 1. Fixed concentrations of a protein and a lipid vesicle in 200 μL of the assay buffer (20 mM Tris–HCl buffer, pH 7.4, containing 0.16 M NaCl) are added to each well of the plate. Typically, the protein concentration is 100 nM and the lipid vesicle concentration is adjusted to give  80% of maximal FP quenching. 2. A fixed concentration of different inhibitors is added to each well. The volume of dimethyl sulfoxide (DMSO; ThermoFisher, Waltham, MA, USA, Cat. No. D128-500) (i.e., 2%, v/v) is minimized to avoid protein denaturation. The inhibitor concentration is typically set at 20 μM

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for initial screening and is gradually lowered for the following rounds of screening to increase the detection threshold. 3. For background correction for each row, the assay is run with the mixture minus lipid vesicles in the next row. 4. The plate is incubated at 25°C with extremely gentle shaking for 10 min. 5. The increase in EGFP fluorescence intensity at 509 nm is monitored with excitation set at 488 nm. 2.7.2 Determination of Inhibition Constant 1. Fixed concentrations of a protein and a lipid vesicle in 200 μL of the assay buffer are added to each well of the plate. Typically, the protein concentration is 100 nM and the lipid vesicle concentration is adjusted to give  80% of maximal FP quenching. 2. An increasing concentration of an inhibitor is added to each well of a given row. The total volume of DMSO (i.e., 2%, v/v) is kept constant for all wells. 3. The plate is incubated at 25°C with extremely gentle shaking for 10 min. 4. The increase in EGFP fluorescence intensity at 509 nm is monitored with excitation set at 488 nm. 5. For background correction for each row, the assay is run with the mixture minus lipid vesicles in the next row.

3. DATA ANALYSIS Since all lipid molecules are associated with membranes, protein–lipid interaction is a complex process that involves membrane adsorption of protein molecules, lateral diffusion of membrane-bound proteins and lipids, binding of a lipid(s) into a lipid pocket, reorientation and/or aggregation of membrane-bound proteins, membrane penetration of proteins, etc. (Cho & Stahelin, 2005). Thus, unlike protein–ligand binding that can be described in a straightforward manner, protein–lipid binding must be analyzed using simplified models. The simplest model for membrane binding is to assume that each protein (P) binds independently to a site on the membrane composed of n lipids (L) (i.e., [P] + [L]/n $ complex) with the dissociation constant as Kd ¼ [P]free [free sites on membrane]/[occupied site on membrane]. Then, binding can be analyzed by a Langmuir adsorption isotherm using a quadratic equation (Eq. 1; Cho et al., 2001):

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½P 0 + Kd + ½L 0 =n 

qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ffi 2 ½P 0 + Kd + ½L 0 =n  4½P 0 ½L 0 =n 2½P 0 (1)

where [L]0, [P]0, and [P]b are total lipid, total protein, and bound protein concentrations, respectively. In our fluorometric assay, ΔF/ΔFmax ¼ [P]b/[P]0, where ΔF and ΔFmax are the fluorescence quenching and the maximal quenching, respectively. It should be noted that this simplified model works successfully when the surface coverage of proteins is low enough to avoid a significant degree of protein–protein interaction. Nonlinear least-squares analysis of either the ΔF/ΔFmax vs [L]0 plot with a fixed value of [P]0 or the ΔF/ΔFmax vs [P]0 plot with a fixed value of [L]0 using Eq. (1) will yield values of Kd and n. The two plots are obtained by titrating lipid vesicles into the protein solution and by titrating a protein into lipids, respectively. The selection of the two approaches depends on several experimental factors, including the extent of the background signal. Finally, it should be noted that since Kd is expressed in terms of molarity of protein-binding sites on the membrane, nKd is the dissociation constant in terms of molarity of lipid monomers. If two related proteins (e.g., wild type and a mutant) bind the vesicles in essentially the same modes, their n values should be comparable and their relative membrane affinity can be assessed from the ratio of Kd values. Otherwise, the ratio of nKd values should indicate the relative affinity, as the nKd to [L]0 ratio, but not the Kd to [L]0 ratio, determines the relative population of membrane-bound protein (¼[P]b/[P]0) at a given [L]0 (e.g., Eq. 1 becomes [P]b/[P]0 ¼ 1/(1 + nKd/[P]0) when [L]0 > n[P]0). It should be stressed that the above quantitative treatment is only an approximation, as the protein-binding site on the membrane has no permanent reality, and therefore binding parameters should be interpreted with caution. Furthermore, binding of a protein to heterogeneous lipid mixtures, including biological membranes, could be much more complex. More advanced isotherms for membrane binding of proteins have been reported (Heimburg, 2007), but they are not practical for the large-scale and highthroughput analysis of membrane-binding data. For those proteins whose membrane-binding isotherms cannot be explained by a Langmuir isotherm, apparent dissociation constant ((Kd)app) is defined as the lipid concentration giving rise to half-maximal membrane binding of protein and is estimated graphically from the binding curve. When a protein selectively binds a specific lipid (e.g., PtdInsP) in the membrane, interfacial dissociation constant

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(Ks) for the lipid (s) can be determined by measuring Kd (or (Kd)app) values at different mol% of s (Xs) (11). Curve fitting of the 1/(Kd) vs Xs plot to an equation, 1/(Kd) ¼ (1 + Xs/Ks)/(Kd)0, where (Kd)0 is Kd in the absence of s, yields the value of Ks.

3.1 Specificity Data Analysis 1. The background fluorescence values are subtracted from binding data. 2. Resulting membrane-binding data of proteins are analyzed using the equation: ΔF/ΔFmax ¼ 1/(1 + [L]%/K). [L]% and K are mole% of a particular lipid and the [L]% value causing half-maximal quenching, respectively. 3. Lipid specificity of a protein can be expressed in terms of relative K values. For example, if the protein is specific for PtdIns(3,4,5)P3, K for PtdIns(3,4,5)P3 should be much (e.g., >5-fold) lower than those for other PtdInsPs.

3.2 Binding Data Analysis 1. The background fluorescence values are subtracted from binding data. 2. Values of n and Kd are determined by nonlinear least-squares analysis of the ΔF/ΔFmax vs [P]0 or ΔF/ΔFmax vs [L]0 plot using Eq. (1). 3. Ks for a specific lipid is determined by nonlinear least-squares analysis of the 1/(Kd) vs Xs plot using the equation, 1/(Kd) ¼ (1 + Xs/Ks)/(Kd)0 (see earlier).

3.3 Inhibition Data Analysis 1. The background fluorescence values are subtracted from inhibition data. 2. Inhibition of membrane binding of a protein by an inhibitor is analyzed using an equation: ΔF ¼ ΔF0/(1 + [I]/Ki) (Dua & Cho, 1994). ΔF and ΔF0 indicate the fluorescence intensity decrease of EGFP by dabsylPE-containing vesicles in the presence and the absence of a given concentration of inhibitor, respectively. [I] and Ki are the free inhibitor concentration and the inhibition constant.

ACKNOWLEDGMENTS The work is supported by National Institutes of Health Grants GM68849 and GM110128.

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