An ELISA-Based Screening Platform for Ligand–Receptor Discovery

CHAPTER THIRTEEN

An ELISA-Based Screening Platform for Ligand–Receptor Discovery Sinem Ozgul*, Sventja von Daake*, Sumie Kakehi*, Davide Sereni*, Natalia Denissova†, Carlie Hanlon†, Yuanpeng Janet Huang†, John K. Everett†, Cuifeng Yin†, Gaetano T. Montelione†,‡,§,1, Davide Comoletti*,¶,||,1,2 *Child Health Institute of New Jersey, Robert Wood Johnson Medical School, Rutgers, The State University of New Jersey, New Brunswick, NJ, United States † Center for Advanced Biotechnology and Medicine, Rutgers, The State University of New Jersey, Piscataway, NJ, United States ‡ Department of Molecular Biology and Biochemistry, Rutgers, The State University of New Jersey, Piscataway, NJ, United States § Department of Biochemistry and Molecular Biology, Robert Wood Johnson Medical School, Rutgers, The State University of New Jersey, Piscataway, NJ, United States ¶ Department of Neuroscience and Cell Biology, Robert Wood Johnson Medical School, Rutgers, The State University of New Jersey, New Brunswick, NJ, United States jj Department of Pediatrics, Robert Wood Johnson Medical School, Rutgers, The State University of New Jersey, New Brunswick, NJ, United States 1 Corresponding author: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. Overview of Workflow 3. The High-Throughput Screen 3.1 Target Selection, DNA Synthesis, and Construct Design 3.2 HEK293 Mammalian Expression System 3.3 Expression of Ectodomains and Secreted Proteins 3.4 PPI ELISA 3.5 Detailed Protocol for 384-Well Plate PPI ELISA 4. Data Analysis and Laboratory Information Management 5. Validation of PPIs and Biophysical Characterization 5.1 Binding Assays 5.2 NMR Assays 5.3 Small-Angle X-Ray Scattering and X-Ray Crystallography 6. PPI ELISA Results

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Current Address: School of Biological Sciences Victoria University of Wellington, Wellington, New Zealand.

Methods in Enzymology, Volume 615 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2018.10.001

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2019 Elsevier Inc. All rights reserved.

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7. Comparison With Affinity Chromatography Methods 8. Future Prospects Acknowledgments References

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Abstract Cell surface molecules are important for development and function of multicellular organisms. Although several methods are available to identify ligand–receptor pairs, ELISA-based methods are particularly amenable to high-throughput screens. ELISA-based methods have high sensitivity and low false-positive rates for detecting protein–protein interaction (PPI) complexes. Here, we provide a detailed protocol for a 384-well ELISA-based PPI screening protocol for the identification of novel cell surface ligand–receptor interactions, together with considerations for validation of PPIs by biophysical methods. This PPI screen has been developed and tested for discovery of novel ligand–receptor pairs between human synaptic adhesion proteins, believed to play crucial roles in many steps of neurodevelopment, from neuronal maturation, to axon guidance, synapse connectivity, and pruning.

ABBREVIATIONS τc AP BLI ELISA FOB HEK HEPES HRP HSQC ITC NMR PBS PEI PNPP PPI PTMs SAXS SPR TMB

molecular rotational correlation time alkaline phosphatase biolayer interferometry enzyme-linked immunosorbent assay fold over background human embryonic kidney cells 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid buffer horse radish peroxidase heteronuclear single-quantum coherence NMR spectroscopy isothermal titration calorimetry nuclear magnetic resonance spectroscopy phosphate buffered saline polyethylenimine para-nitrophenylphosphate, substrate of AP protein–protein interaction posttranslational modifications small-angle X-ray scattering surface plasmon resonance protein interaction detection 3,30 ,5,50 -tetramethylbenzidine, an immunohistochemistry stain

1. INTRODUCTION The completion of the human genome, together with outstanding technical developments in proteomics and laboratory automation, now provides the potential for mapping protein–protein interactions (PPIs) at an unparalleled speed and scale. These advances will enable the generation of

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comprehensive and accurate PPI maps for both intracellular and extracellular proteins. For example, the cell surface protein atlas (Bausch-Fluck et al., 2015) provides a mass spectrometry-derived cell surface protein atlas; comprehensively called surfaceome. This collection reports experimentally determined glycosylated cell surface proteins from 72 different cell types, providing a combined dataset of 1492 human and 1296 mouse cell surface proteins. Using a large collection of public databases and manual literature searches Ramilowski et al. (2015) have compiled a list of 1894 known ligand–receptor pairs. This list includes ligand–receptor pairs from 144 cell types, encompassing neuronal, endothelial, hematopoietic, epithelial, and mesenchymal cells. Classic affinity chromatography coupled with mass spectrometry has been successfully used for decades for ligand–receptor discovery (Olsen, Meunier, & Changeux, 1972; Savas et al., 2014). More recently, other systematic approaches to discover new ligand–receptor pairs, such as Tandem affinity purification (TAP) tagging (Puig et al., 2001), LUMIER (luminescence-based mammalian interactome) (Barrios-Rodiles et al., 2005), MAPPIT (mammalian protein interaction trap) (Eyckerman et al., 2001), and other methods (Kang et al., 2014; Linhoff et al., 2009), have been successfully implemented in the discovery of new cell surface receptors or to deorphanize known cell surface receptors. None of the methods, however, are easily amenable for high-throughput implementation. In the past decade, taking advantage of variations of the classic enzymelinked immunosorbent assay (ELISA) (Engvall & Perlmann, 1971), several groups have described the discovery of dozens of new ligand–receptor protein pairs from Drosophila melanogaster, Zebrafish, and a variety of mammalian proteins (Kerr & Wright, 2012; Ozkan et al., 2013; Visser et al., 2015; Wojtowicz et al., 2007). These PPI ELISA-based assays have the potential for high-throughput screening of hundreds, or thousands, of potential protein interaction partner pairs. However, the currently published protocols have been performed manually in 96-well plate format. In this monograph, we describe an improved version of these PPI ELISA-based protocols for secreted proteins and/or ectodomains of transmembrane proteins, which has been adapted to a 384-well format and uses human neuronal proteins that are expressed using suspension-adapted human embryonic kidney (HEK) 293F cells in serum-free medium. Our current primary application of PPI ELISA technology is directed to human secreted proteins, and ectodomains of transmembrane proteins, associated with the neuronal synapse. In the mammalian brain, the specification of each synaptic contact and the precise and dynamic balance between excitatory and inhibitory connections are enabled by a rich repertoire of cell

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adhesion and signaling molecules with diverse structural and functional identities (de Wit & Ghosh, 2016; O’Rourke, Weiler, Micheva, & Smith, 2012). Despite decades of intense research, however, the majority of neuronal and glial cell surface proteins do not have a known bone fide binding partners. Protein interaction functions of these proteins are often modulated by various posttranslational modifications (PTMs), including disulfide bond formation and glycosylation, requiring the use of mammalian expression hosts for protein production.

2. OVERVIEW OF WORKFLOW The overall workflow of this protocol begins with bioinformatics studies to identify the genes of interest that will be tested for PPIs. Because of the free access to numerous public protein and gene databases, in addition to libraries already published, it is usually straightforward to identify or build the appropriate lists of protein interaction candidates. In addition, as these databases (e.g., Uniprot.org) are generally carefully curated, the architecture of the proteins (e.g., domain organization and interdomain boundaries) necessary to derive cloning boundary information is usually correct. Once the genes of interest have been cloned or synthesized, and amplified (e.g., as mini- or midi-preps), the amount of protein present in cell cultureconditioned medium after transient protein expression is normally adequate for downstream experiments. The ELISA-based PPI assay itself is very rapid. It is a standard ELISA format, modified to provide a “sandwich” assay (Wojtowicz et al., 2007), in which the complex to be detected, a Fc fusion protein bound to an alkaline phosphatase (AP) fusion protein, is bound between two antibodies—the capture antibody and the detection antibody (Fig. 1). We have adopted the VIAFLO 96/384 (Integra) 96- and 384channel pipette and Tecan HydroSpeed 384-well microplate washer, together with liquid-handling robotics, to provide a 384-well PPI ELISA protocol. Using these liquid handlers and proper organization, two fulltime-equivalent scientists can run 25–30 384-well plates in a single day (9600 to 11,500 binary interactions in a day), and easily run four experiments in a week (45,000 binary interactions in a week). Additional time and throughput considerations are outlined in Table 1. Data interpretation is usually fast as the true- and false-positive wells are easily spotted when multiple plates are run during the same session.

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Fig. 1 Schematic representation of the two plasmids used in this protocol and the ELISA orientation. (A) The Fc fusion protein contains a leader peptide (PLP, prolactin leader peptide) followed by a FLAG tag N-terminal to the gene of interest. The gene, cloned by NotI–XbaI, is followed by a 3CPro cleavage site and the dimeric human Fc domain. Similar to Fc construct, the AP fusion protein contains a leader peptide and a FLAG tag N-terminal to the gene of interest. C-terminal to the gene, cloned by NotI–XbaI, there is a human AP gene followed by a His6 tag. (B) Schematic of the ELISA orientation showing tetramerization of both bait and prey driven by the antibodies. (C) LINGO2-AP was used to coat every well of the plate (bait) and a panel of 320 different proteins was added to the wells. Few nonspecific (or false-positive) interactions, visible in every plate run the same day, were detected (red squares). (D) Using EphrinA3-AP as bait, the same panel of 320 preys was added. In addition to the nonspecific interactions of the LINGO-2 plate, Eph-A 1–8 receptors were, as expected, unequivocally identified (Gale et al., 1996).

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Table 1 Time Line for High-Throughput PPI ELISA Screen Activity Time to Complete

Gene design

Days to weeks

Commercial gene synthesis

4–6 weeks depending on the size of the library

Gene amplification (mini- or midiprep) Typically weeks, but it depends on the size of the library Transient protein expression

1 week

Testing of protein expression (e.g., Western blot)

1–3 days, depending of the number of samples to test

ELISA

2–3 days depending on the number of plates to run

Data interpretation

1 day per set of experiments (e.g., 30  384-well plates)

Data validation (ITC, BLI, SPR, NMR, SAXS, X-ray crystallography, etc.)

Weeks to months depending on the number of positive interactions to test and the technology employed

3. THE HIGH-THROUGHPUT SCREEN 3.1 Target Selection, DNA Synthesis, and Construct Design The initial component of the screen is the selection of target proteins for PPI analysis. To select synaptic protein ectodomain sequences of interest, we used the Universal Protein Resource (UniProt) database (Uniprot.org). UniProt is a comprehensive resource for protein sequences and a variety of other annotations. This information was supplemented with data on human proteins and their interaction partners curated from the literature and other databases, and organized in our Human Cancer Protein Interaction Network (HCPIN) database (Huang et al., 2008). In the absence of other data, the extracellular domain (ectodomain) of each target protein was defined as the region starting from the first amino acid after the signal peptide to the last amino acid before the transmembrane domain, the ω-site for Glycosylphosphatidylinositol (GPI)-anchored proteins (Eisenhaber, Bork, & Eisenhaber, 1998), or the very last amino acid of secreted proteins. In many cases, protein construct design may be further guided using signal peptide, secondary structure, and disorder predictions available from the

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DisMeta Protein Construct Design server (Huang, Acton, & Montelione, 2014), as we have described elsewhere (Acton et al., 2011). Genes are custom synthesized with human codon optimization and cloned into expression vectors. Typically, >90% of the genes we tested were secreted by HEK293 cells at levels compatible with downstream experiments, confirming that the cloning boundaries are correct. In our library, many of the nonexpressing genes turned out to be hetero-oligomers (e.g., GABA B1–2), which need to be cotransfected with another gene in order to provide useful levels of protein expression. DNA coding for target genes is next cloned in frame with either a C-terminal Fc or an AP tag (Fig. 1A). These expression tags enhance protein production, solubility, and enable detection and purification of the secreted proteins. Both constructs also carry a DYKDDDDK tag at the N-terminus of the protein, inserted after the signal peptide. A distinctive advantage of the DYKDDDDK tag is that it can be cleaved, as the DDDK sequence is an enterokinase (EK) cleavage site, leaving an N-terminal Asp residue. Between the end of the target protein of interest and the beginning of Fc domain, there is also a human rhinovirus HRV3C protease (3CPro) cleavage site. The presence in the Fc vector of two different cleavable tags at N- and C-terminal ends enables flexible purification strategies useful to rapidly move into structural biology studies. The AP construct differs from the Fc construct because it lacks the 3CPro cleavage site, contains AP as a C-terminal tag, and carries a hexaHistidine (His6) tag at its C-terminus, in frame with the AP. The fact that AP is enzymatically active provides a simple way to test expression and protein concentration. Importantly, both of these tags are dimeric, a property that increases the avidity of the ligand–receptor interaction and effectively compensates for the low affinity of some of the potential interactions.

3.2 HEK293 Mammalian Expression System Secreted or extracellular domains of transmembrane proteins are subjected to two main types of PTMs: disulfide bonding and O- and N-linked glycosylation (Walsh, 2005). These PTMs are crucial for proper folding of the protein, trafficking and secretion, and for activity. Therefore, mammalian expression systems such as HEK293 or Chinese hamster ovary (CHO) cells are the method of choice to guarantee that the recombinant protein fragment is folded properly and retains its binding properties (Comoletti et al., 2003; Ranaivoson et al., 2015; Rubio-Marrero et al., 2016). Another key advantage of this expression system is the inherent quality control of the

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folding. If folding does not occur or it remains incomplete (e.g., single point mutations, incorrect cloning boundaries), the endoplasmic reticulum chaperones retain the nascent protein and secretion is inhibited. In practical terms, a protein that is not optimally folded is not secreted (De Jaco et al., 2010) and hence it will not be used in the screen. We have adopted the use of high-density suspension-adapted HEK293F cells (available from ThermoFisher). These cells are easy to maintain in culture (e.g., they have a fast growth rate, splitting is done by dilution, FreeStyle™ and other media maintain growth and expression for several days after transfection) and they generally provide excellent expression of secreted proteins. All of these advantages also help reduce labor and costs associated with the generation of classical stable adherent cell lines. Moreover, as these cells grow in serum-free medium, no additional proteins (e.g., immunoglobulins) are introduced into the ELISA-based assay. Typically, HEK293F cells are maintained in FreeStyle™ 293 Expression Medium on a platform shaker in a humidified incubator at 5% (v/v) CO2 and 95% (v/v) air, and kept for 20–25 generations before being replaced with a new vial to avoid genetic drifting and minimize mycoplasma contaminations. Cells should be maintained between 5  105 and 4  106 cells/mL in a volume 10%–15% the total volume of the culture flask, so that thorough mixing and oxygenation are achieved. Correct rotation rate of the platform should be determined for each flask type and culture volume (Hacker et al., 2013).

3.3 Expression of Ectodomains and Secreted Proteins HEK293F cells are readily transfected with polyethylenimine (PEI), which is very economical and easy to use. PEI is normally prepared as a stock solution at 1 mg/mL in PBS or similar buffer (e.g., 20 mM HEPES, pH 7.5, and 150 mM NaCl). To dissolve PEI, 100 mg of the powder is added to the appropriate volume of warm PBS and stirred vigorously on a hot plate set to 75°C until completely dissolved. Once dissolved and cooled to room temperature, the solution is sterile filtered by using a 0.22-μm syringe filter, aliquoted (e.g., 1 mL or other size, depending on the downstream use), and frozen at 80°C until needed. On the day of transfection, cells are counted with a hemocytometer, and the required number of cells (e.g., 0.7  106 for Fc-ectodomain constructs, see below) are centrifuged at 125  g for 5 min. The cells are then resuspended in fresh FreeStyle™ medium at 1  106 cells/mL. The

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viability of cells should be >95%. Fresh 293F FreeStyle™ medium is critical for high transfection efficiency. We have compared transfection efficiency of enhanced green fluorescent (eGFP) protein by using FreeStyle (Thermo), EX-CELL® 293 (Sigma), or Pro293™ (Lonza) media, and we found that FreeStyle™ has about twice as many transfected cells than Pro293™, whereas EX-CELL 293 media show no significant signs of transfection (data not shown). Once the cells have been thoroughly resuspended in the new medium, they are placed back into the incubator until needed. When transfecting hundreds of Fc-ectodomain constructs in small volumes (0.7 mL), transfections are performed in 24-well plates. While the freshly resuspended cells are in the incubator, the appropriate plasmid cDNA (2 μg/million cells) and PEI (6 μL/million cells) are added to each well. To reduce error in the cDNA delivery, all our Fc-tagged plasmids are prepared and stored at a standard concentration of 0.2 μg/μL. At this point, the 24-well plates in which DNA, PEI, and cells have been added are placed in the incubator for expression over the next 4–5 days. For the AP constructs we use 4 mL of medium at 1  106 cells/mL, and transfect with proportional amounts of plasmid cDNA and PEI in a 1:3 ratio (DNA/PEI). Transfections are performed in 50-mL conical tubes with ventilated caps. Finally, valproic acid (VPA) can be added to the cell culture to a final concentration of 2.2 mM to boost protein expression (Hacker et al., 2013). A stock solution of 220mM VPA in water allows for the addition of 10 μL of VPA per 1 mL of final transfection volume. At 5 days posttransfection, the cell culture medium is harvested by centrifugation and the cells discarded. The Fc-ectodomain media are carefully transferred with a multichannel into a final 96-well plate and stored at 4°C for immediate use or up to 1 week. Fc fusion proteins are quantified and quality checked by Western blot, using known quantity of Fc-only protein as a standard, and either diluted or concentrated to a final concentration of 10–20 ng/μL. AP fusion proteins are quantified using a para-nitrophenylphosphate (PNPP) assay, by comparing the activity of each protein to known amount of calf intestinal alkaline phosphatase (CIP, New England Biolabs) as a standard, and typically diluted to 40 pg/μL.

3.4 PPI ELISA Immobilization of the AP protein (antigen) is accomplished indirectly via a capture antibody (anti-AP) that has been adsorbed to the microtiter plate. The remaining unbound surface area is then blocked by a solution of an

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irrelevant protein (1% casein or 5% dry nonfat milk) to prevent other proteins (including the detection antibody) from adsorbing to the plate during subsequent steps. The ideal blocking buffer will coat the plastic well, without altering or obscuring the epitope for the interactions, while not binding other bait proteins. Conditioned media containing the appropriate ectoFc (first) and ecto-AP (second) proteins are added sequentially to the plate followed by anti-Fc antibody, which helps clustering the ecto-Fc prey by increasing the apparent affinity of the potential interaction. After 4 h, the plates are washed, 1-Step Ultra TMB-ELISA horseradish peroxidase (HRP) substrate (see Reagents) is added, and the presence of bound Fc prey detected by the development of a blue color that can be read at 650 nm by a Spectramax i3 plate reader (Fig. 2). Positive controls, known PPI pairs (e.g., βNRXN/NLGN1 and FLRT3/LPHN3), are used to gauge the sensitivity of the assay in each plate and negative controls (e.g., wells where the AP antibody is omitted or where the Fc-only sample is added) are necessary to obtain background values used to quantify the positive reactions. With this system, we could detect interactions with affinities of 10 μM or better (Ranaivoson et al., 2019). As an example, positive controls βNRXN/ NLGN1 have measured binding affinity of 100 nM (Comoletti et al., 2006) and FLRT3/LPHN3 has measured affinity of 20 nM (Ranaivoson et al., 2019), but we rarely see the interaction of CASPR2/ CNTN1, which has an estimated affinity of 20 μM (Rubio-Marrero et al., 2016).

3.5 Detailed Protocol for 384-Well Plate PPI ELISA In the following sections, we provide our standard operating protocol for the 384-well PPI ELISA. Experiment overview • Coating of the plate with anti-AP antibody, incubation overnight in cold room. • Wash plate (washing buffer 1). • Block wells with 1% casein or 5% dry nonfat milk. • Wash plate (washing buffer 2). • Add conditioned medium containing ecto-Fc. • Add conditioned medium containing ecto-AP mixed with anti-Fc-HRP antibody. • Incubate 4 h at room temperature. • Wash plate (washing buffer 2).

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Fig. 2 Blocking reagents. Plates blocked with different blocking reagents display diverse false-positive interactions. In our conditions and with our library of prey, the best blocking reagent was 5% dry milk. The other two blocking agents display the same false-positive wells (red squares), in addition to several others in different position.

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Add the TMB substrate and read plate at 650 nm. Scan with an office scanner to obtain an image of the plate to match with the ABS reading. Empty and wash with PBS, add the PNPP, 1 h incubation. Read at 405 nm and scan to determine the amount of ecto-AP protein immobilized to the plate.

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Reagents and supplies • Nunc-Immuno MaxiSorp 96-well plates; Nunc; Cat # 442404. • Plate sealers; Fisher Scientific; Cat # 1256790. • PLAP Ab-1 (8B6.18) Mouse Mab; 1 mg/mL; NeoMarkers; Cat # MS-208-PABX. • 5% Alkali-soluble casein; Novagen; Cat # 70955-225ML. • HEK293F; ThermoFisher; Cat # R790-07. • FreeStyle™ 293 Expression Medium; Cat # 12338018. • PEI (25 kDa linear PEI; Polysciences, Inc.; Cat # 23966). • VPA (Sigma; Cat # P4543-100G). • Ectodomain-AP conditioned culture medium. • Ectodomain-Fc conditioned culture medium. • Mouse anti-human Fc-HRP antibody 1 mg/mL; Serotec; Cat # MCA514P. • TMB substrate; 1-Step Ultra TMB-ELISA; Thermo Scientific; Cat # 34028. • Washing buffer 1 (WB1): 1  PBS-Tween (Tween ¼ 0.01%). • Washing buffer 2 (WB2): HEPES 20 mM, NaCl 150 mM, MgCl2 2 mM, CaCl2 2 mM. • VIAFLO ASSIST (Integra), automated electronic 16-channel pipette. • VIAFLO 96/384 (Integra), electronic 96- and 384-channel pipette that enables fast and precise transfers of up to 384 samples to microplates. • Tecan HydroSpeed, 384-well microplate washer. Experimental procedure (volumes for one 384-well plate testing one AP-tagged protein against a library of 382 different Fc-tagged proteins). Day 1: Coating of microtiter plates • Dilute the anti-AP antibody in 1  PBS to 3 μg/mL and add 15 μL in each well by using the VIAFLO ASSIST. • Seal the plate with the pleat sealer. • Spin the plate (1000  g for 1 min) to ensure solutions are at the bottom of the plate and no air bubbles are trapped. Visually inspect. • Incubate overnight in the cold room (4°C). • Prepare the casein dilution (1% in 1 PBS) or dry milk (5% in 1  PBS) for the next day. Day 2: Perform ELISA • Remove the anti-AP antibody with the plate washer and wash twice with WB1. Immediately add 100 μL of 1% casein in each well to block the plastic using the VIAFLO ASSIST. • Spin (1000  g for 1 min) and incubate plates containing 1% casein 1 h at room temperature.

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Manually remove the casein by quickly inverting the plate above the sink and blotting on paper towel. Add 7 μL of a solution made by mixing 5 μL of ecto-AP (same ecto-AP protein for each well) and 2 μL of HRP anti-Fc (1:80 in 1% casein in 1  PBS) using the VIAFLO ASSIST. To prepare the solution, first dilute the HRP anti-Fc Ab in 1% casein in PBS then add the total amount of antibody solution to the total amount of ecto-AP protein. For example, for each 384-well plate, 768 μL of diluted anti-Fc antibody is added to 1920 μL (5 μL  384) of ecto-AP medium. This protocol will minimize the error of delivering 2 μL several times in each well. (Note— Increase volumes by 10% to account for void volumes of the pipetting robot.) Add 5 μL of ecto-Fc protein, by using the VIAFLO 384 from four 96-well plates to a single 384-well plate, in four steps. Control wells: Designate two wells for the positive and negative controls, and manually pipet into those wells the corresponding control proteins. Seal the plate with the plate sealer. Spin the plate (1000  g for 1 min) Incubate at room temperature for 4 h. Remove the content of the plate with the plate washer and wash three times with WB2. Add 30 μL of TMB substrate by using the VIAFLO ASSIST. Incubate the plates at room temperature for 1 h before reading. A blue color should develop within minutes in the positive control well and the positive wells. Scan the plate at 650 nm with a Spectramax i3 plate reader or similar instrument. Scan the plates with a picture scanner to have the equivalent picture of the plate with blue wells (see, for example, Figs. 1C and 2). Wash the plate twice with PBS using the plate washer. Add 15 μL of PNPP substrate to each well using the VIAFLO ASSIST. Incubate 1 h by keeping the plate in the dark. Scan and read the plate at 405 nm to measure the amount of ecto-AP immobilized to the plate. Results of the 650 nm scans are quantified by calculating the background OD650 value of the negative control wells (or other well that remained clear) and then dividing the OD650 value for such background.

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4. DATA ANALYSIS AND LABORATORY INFORMATION MANAGEMENT Because we measure colorimetric values up to 20-fold over background (FOB), we have adopted the convention of defining positive interaction as wells that show an OD650 with colorimetric values >3–4 FOB, and that are present in only one plate in a given experiment. True positives were also validated by observing the interaction also in the reverse orientation of prey and bait; i.e., if interaction A–B is observed in the Fc-A + B-AP well, it is validated by its detection also in the Fc-B + A-AP well. Certain preys appear to bind specifically to the protein used as blocking reagent (Fig. 2), representing false-positive readings. We tested three different blocking reagents side by side (5% dry milk, 5% fetal bovine serum, and 1% casein), and observed that under our experimental conditions, 5% dry milk has the best blocking activity, as it results in fewest false positives. The identification of false-positive interactions relies on the signal generated by the blocked plate and by running multiple plates in a single session. In these conditions, false-positive interactions are readily detected and excluded from further examination, whereas real hits can be productively pursued. Proteins with high isoelectric point (pI) > 9.0, or long stretches of positively charged amino acids, were sometimes observed to give falsepositive results. These basic proteins tend to bind nonspecifically to the blocking agent, casein, which has a pI  4.5 (Ranaivoson et al., 2019). Probably because of the steric hindrance of the tag with the binding site, occasionally only one orientation leads to a positive interaction. Therefore, the subsequent bibliographical search (for previously observed interactors) and/or further biophysical validation of the PPI is required to distinguish between true- and false-positive interactions. Although false-negative rates are difficult to estimate, Ozkan et al. (2013) found that they could identify all the positive interactions that were tested in the system. This PPI ELISA generates an extensive amount of data and reagents, which must be intelligently organized and tracked. Our laboratory information management system, called SPiNE (Goh et al., 2003), was modified for this project to track experimental results (e.g., protein expression, protein purification and 96-well-based ELISA screens, molecules with synaptogenic activities) and products (e.g., expression vectors and individual purified protein samples). SPiNE’s web-based interfaces allow researchers to archive, store, retrieve, and share results between laboratories, and to utilize and contribute to standardized data and material repositories.

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5. VALIDATION OF PPIs AND BIOPHYSICAL CHARACTERIZATION 5.1 Binding Assays To ensure that positive interactions do not arise as an artifact of the ELISA protocol, each novel (i.e., unpublished) interaction must be tested with orthogonal binding assay such as biolayer interferometry (BLI), surface plasmon resonance (SPR), or isothermal titration calorimetry (ITC). An example of validation using these binding assays is reported in Fig. 3 (Ranaivoson et al., 2015; Rubio-Marrero et al., 2016).

5.2 NMR Assays NMR provides alternative approaches for validation of proposed PPI partners. These studies can be done using <50 μg of protein samples in microcryogenic probes (Rossi et al., 2010). However, there are several caveats that limit these NMR experiments. In general, proteins should be isotopeenriched with 15N and 13C, and for proteins larger than about 20 kDa also with 2H, for optimal detection schemes, and also for determining backbone resonance assignments using triple-resonance NMR methods. In our studies, isotope enrichment is done most economically by production of protein samples in E. coli expression hosts (Acton et al., 2011). As the genes for the interacting protein partners are already synthesized for the PPI ELISA, it is generally straightforward to subclone interacting proteins or their domains in E. coli expression hosts. Depending on the nature of the protein partners, it may be necessary to generate protein samples with native disulfide bonds. For this purpose, we have developed an efficient system for production of 13 15 C, N- and 2H,13C,15N-enriched disulfide containing proteins as maltose-binding protein (MBP) fusions expressed in the E. coli BL21 trxB (DE3) cells (Zhang et al., 2016). The E. coli trxB strain lacks thioredoxin reductase, which allows formation of disulfide bonds within the E. coli cytoplasm (Derman, Prinz, Belin, & Beckwith, 1993). Fusion with MBP enhances the expression and solubility of the construct, suppresses aggregation, and generally provides for higher yields of native folded proteins (Kapust & Waugh, 1999). The isotope-enriched proteins of interest are then cleaved from their MBP tags using multiple engineered protease cleavage sites between the MBP and the target proteins (Zhang et al., 2016), providing samples with native disulfide bonds suitable for NMR studies. Such E. coli production systems, however, do not provide for some key PTMs,

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Fig. 3 Positive interaction validation. Following the ELISA identification of new PPIs, different binding assays can be used to validate the interactions and to calculate dissociation constants for each pair. Two examples follow. (A) Biolayer interferometry (BLI) experiment of the association between the purified extracellular domains CNTN1 and CASPR2. Five concentrations of CASPR2 were used to determine the affinity of the association, calculated in the inset. (B) Isothermal titration calorimetry (ITC) characterization of the interaction of LPHN3 and FLRT3. 100 μM of LPHN3 was injected into (Continued)

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such as glycosylation, which may be important in mediating PPIs between secreted proteins. With such samples, PPIs can be detected by changes in rotational correlation times, chemical shifts, or other NMR spectral parameters upon complex formation. We generally utilize 1D 15N T1/T2 measurements to measure changes in rotational correlation times τc that accompany complex formation (Acton et al., 2011; Aiyer et al., 2014; Aramini et al., 2011, 2015; Rossi et al., 2010) and [15N–1H]-TROSY HSQC spectra (Pervushin, Riek, Wider, & Wuthrich, 1997) to detect changes in chemical shifts resulting from complex formation. Full structure determination by NMR provides details of the protein–protein complex, but is generally limited to complexes with molecular weights <50 kDa. Accordingly, these extensive structural studies of larger protein complexes may require trimming the interaction partners down to the specific subregions which mediate the PPI.

5.3 Small-Angle X-Ray Scattering and X-Ray Crystallography In addition to NMR spectroscopy, protein complexes of particular interest detected in the PPI ELISA screen can be further investigated by other structural biology techniques. X-ray crystallography and small-angle X-ray scattering (SAXS) do not require isotope enrichment, and can be done on proteins produced in HEK293 expression systems. They are often used in tandem because they offer complementary information that strengthens the interpretation of the data. For example, whereas crystallography provides high-resolution structural data, in dealing with complexes of two or more proteins one needs to rule out artifacts due to crystal lattice packing. SAXS, on the other hand, is a lower resolution technique but can be performed in solutions of nearly physiological buffers. SAXS is particularly valuable for detecting complex formation and for determining the oligomeric states of complexes (Grant et al., 2011; Putnam, Hammel, Hura, & Tainer, 2007; Trewhella, 2016). Fig. 3—Cont’d the cell containing FLRT3-LRR at 10 μM. Panel (A): Adapted from RubioMarrero, E. N., Vincelli, G., Jeffries, C. M., Shaikh, T. R., Pakos, I. S., Ranaivoson, F. M., et al. (2016). Structural characterization of the extracellular domain of CASPR2 and insights into its association with the novel ligand contactin1. The Journal of Biological Chemistry, 291, 5788–5802. doi:10.1074/jbc.M115.705681. Panel (B): Adapted from Ranaivoson, F. M., Liu, Q., Martini, F., Bergami, F., von Daake, S., Li, S., et al. (2015). Structural and mechanistic insights into the latrophilin3-FLRT3 complex that mediates glutamatergic synapse development. Structure, 23, 1665–1677. doi:10.1016/j.str.2015.06.022.

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6. PPI ELISA RESULTS The high-throughput PPI ELISA has been tested on human proteins associated with neuronal synapses. 205 Genes coding for synaptic proteins (or their domains) were synthesized, and each protein produced in HEK293 cells with Fc and AP different affinity tags. We screened 36,490 interaction experiments using the ELISA-based protein interaction screen in 384-well microtiter plates. The screen identified 197 interactions, 107 of which had been previously reported and are essentially positive controls, and 90 of which were not previously described (Ranaivoson et al., 2019). Among these interactors, we observed that IgLONs, a family of proteins implicated in various human disorders, interact as homo- and heterodimers. These interactions were then validated by BLI, and by determining highresolution X-ray crystal structures of three complexes (Ranaivoson et al., 2019).

7. COMPARISON WITH AFFINITY CHROMATOGRAPHY METHODS To map ligand–receptor pairs of cell surface neuronal proteins, modern affinity chromatography coupled with mass spectrometry-based approaches has been used for decades and is still a reliable way to find novel interacting protein pairs (de Wit et al., 2013; Savas et al., 2015). These approaches, however, also have shortcomings. First, the choice of “bait” is biased by the investigator’s hypothesis or the lab interest. Second, mass spectrometry is so sensitive that using brain lysate produces hundreds of false positives, which need to be scored with some form of statistical analysis. Third, potential candidate proteins have to be validated experimentally, by acquiring and cloning the potential gene targets, and performing confirmatory in vitro binary binding assays. Finally, both low- and very highaffinity interactions are missed because the former dissociate quickly during the relatively long and stringent washing steps, and the latter may not be identified when the native complex remains unavailable to the bait during the “fishing” experiment. ELISA approaches, on the other hand, offer several key advantages. First, although ELISA-based assays are not novel technologies, they are extremely robust techniques. Second, as all extracellular receptors are expressed recombinantly, low abundance or developmentally regulated genes will be present

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in the screen in similar amounts. Third, binary interactions will only reveal direct PPIs; therefore, indirect interactions or cofactors are excluded, dramatically reducing the rate of false-positive interactions typical of other technologies. Fourth, the ELISA screen can identify interactions with dissociation constants ranging from pM up to the 10–20 μM range, which covers a wider range of physiologically relevant interactions. In addition, because the oligomerization of both pray and bait results in signal amplification, even low expression of either protein is rarely a limiting factor and is not uniformly detrimental to detecting interactions (Ozkan et al., 2013). Robotic handling also allows assessment of >10,000 protein interactions per day. Furthermore, once the gene library is completed, it can be rapidly deployed to screen other targets such as entry receptors for neurotropic viruses, and other systems. Limitations of the ELISA are mainly related to the completeness and the cost of the library, and the fact that proteins containing more than one transmembrane domain typically need to be excluded from the screen because no single ectodomain contains the ligand-binding site. Finally, whereas this approach works very well to detect binary interactions, it may not be suitable to detect three (or more)-way complexes, if the multicomplex is obligatory (A + B, A + C, or B + C alone do not bind). However, if two binding partners can nucleate first followed by the third protein (A + B associate first, followed by C to make ABC), then at least the first pair can be discovered, even if the interaction is weaker than in the complete complex.

8. FUTURE PROSPECTS The PPI ELISA described here is a robust, high-throughput approach for identifying interactions between protein pairs. Because the target genes are already cloned in expression plasmids, the reagents provide DNA suitable for subcloning into alternative expression vectors, and ready entry to protein sample production for biophysical validation and follow on structural biology studies. Hence, these studies can create a wide range of opportunities for structural studies of eukaryotic protein complexes by X-ray crystallography, NMR, SAXS, and/or cryoelectron microscopy (cryoEM). The platform described here has been successfully tested for detecting known and novel PPIs between secreted, human synaptic proteins (Ranaivoson et al., 2019). The platform is readily adopted for high-throughput PPI screens of other eukaryotic PPI involving proteins that go through the secretory pathway.

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For NMR studies in which isotope-enriched samples are required, our general strategy is to identify the interaction domains of secreted multidomain constructs, and produce these in E. coli expression systems where isotope enrichment is most cost effective. However, targeted proteins can also be produced with 2H,13C,15N enrichment in certain animal expression systems, such as Pichia pastoris (Zhang et al., 2016, 2017), or in cell-free eukaryotic expression systems (Zhao et al., 2010). Production of isotopeenriched samples in HEK systems is also feasible, though currently quite expensive. In this regard, NMR studies of these complexes need not focus on full structure determination with NMR data, but rather by use of hybrid methods that combine sparse NMR, SAXS, and cryoEM data together with advanced molecular modeling methods.

ACKNOWLEDGMENTS We would like to thank Dr. W. Wojtowicz for generously providing some of the plasmids used in this work, and for priceless advice during the development of the assay. This work was funded in part by National Science Foundation grant MCB-1450895 to G.T.M. and D.C. G.T.M., N.D., and C.H. were supported in part by the Jerome and Lorraine Aresty Endowment Fund. N.D. was also supported in part by a pilot award from the Rutgers Cancer Institute of New Jersey a 2017 Precision Medicine Pilot Award from the Rutgers Cancer Institute of New Jersey (to G.T.M.). This work was also supported in part from NIH Grant R01GM120574 (to G.T.M.), the Robert Wood Johnson Foundation to the Child Health Institute of New Jersey [grant #74260], and 2016 Precision Medicine Pilot Award from the Rutgers Cancer Institute of New Jersey, and the 2017 Busch Biomedical Grant Program to D.C.

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