Protocol for Peptide Synthesis on Spectrally Encoded Beads for MRBLE-pep Assays.

Every living cell relies on signal transduction pathways comprised of protein-protein interactions (PPIs). In many cases, these PPIs are between a folded protein domain and a short linear motif (SLiM) within an unstructured region of a protein. As a result of this small interaction interface (3-10 amino acids), the affinities of SLiM-mediated interactions are typically weak (K ds of ~1-10 µM), allowing physiologically relevant changes in cellular concentrations of either protein partner to dictate changes in occupancy and thereby transmit cellular signals. However, these weak affinities also render detection and quantitative measurement of these interactions challenging and labor intensive. To address this, we recently developed MRBLE-pep, a technology that employs peptide libraries synthesized on spectrally encoded hydrogel beads to allow multiplexed affinity measurements between a protein and many different peptides in parallel. This approach dramatically reduces both the amount of protein and peptide as well as the time required to measure protein-peptide affinities compared to traditional methods. Here, we provide a detailed protocol describing how to: (1) functionalize polyethylene glycol diacrylate (PEG-DA) MRBLE beads with free amine groups, (2) synthesize peptide libraries on functionalized MRBLEs, (3) validate synthesized peptide sequences via MALDI mass spectrometry and quantify evenness of peptide coverage on MRBLEs, (4) use MRBLE-bound peptide libraries in multiplexed protein binding assays, and (5) analyze binding data to determine binding affinities. We anticipate that this protocol should prove useful for other researchers seeking to use MRBLE-pep in their own laboratories as well as for researchers broadly interested in solid-phase peptide synthesis and protein-protein binding assay development.

The recently published MRBLE-pep technology allows the measurement of binding affinities for many peptides in parallel, thereby facilitating protein-SLiM interaction characterization .
In line with one-bead-one-compound libraries, the method synthesizes peptides directly on beads via solid-phase peptide synthesis to generate one-code-one-peptide libraries (Lam et al., 1991  Bead-based assays are extremely useful for multiplexed measurements of protein-peptide affinities and have the potential to directly quantify affinities using small amounts of material. However, successful binding assays depend critically on considerations of reagent concentration, binding equilibria and detection of the binding partners. Moreover, it is essential that binding assays take place at appropriate concentrations. Therefore, high-quality measurements depend critically on a variety of additional technical details, including robust and even functionalization of beads, gentle yet thorough mixing of beads during functionalization and binding assays, careful consideration of peptide density at the bead surface, and the number of beads added to each reaction. Here, we provide a detailed protocol describing best practices for: (1) functionalization of MRBLEs prior to peptide synthesis, (2) solid-phase peptide synthesis directly on functionalized MRBLEs, (3) peptide sequence validation and peptide density measurements, (4) MRBLE-pep protein binding assays, and (5) data analysis. While these results are presented here in the context of the MRBLE-pep assay, we anticipate that this detailed protocol may provide useful information for researchers optimizing any bead-bound binding assay. 3 www.bio-protocol.org/e3669

Procedure
This complete protocol describes ( Figure 1): (1) Functionalization of PEG-DA hydrogel beads in preparation for peptide synthesis, (2) solid-phase Fmoc peptide synthesis on PEG-DA beads, (3) peptide validation and peptide density measurements, (4) multiplexed protein-peptide binding assays using bead-bound peptide libraries, and (5) final data analysis to extract quantitative binding affinity information (Kd). In this protocol, all amounts are calculated for 48 tubes containing one code each (a 48-code peptide library) with an estimated 1 mg of MRBLEs for each code (about 10,000 beads) and a loading capacity of 0.32 mmol/g. Therefore, 1 equivalent ~50 mg ~0.016 mmol.

Part I: Production of MRBLEs
Prior to the start of the protocol, produce PEG-DA hydrogel beads using either a multilayer PDMS 'bead synthesis' microfluidic device (as described previously [Gerver et al., 2012]) or using syringe pumps and standard droplet generators.
Preparation after bead synthesis: 1. Transfer MRBLEs containing each spectral code (here, a 48-code library) into 48 x 2 ml reactor vessels placed in a U-Block Reactor mounted in a vacuum manifold for easy bead washing without bead loss (Figure 2 and see Notes). For each wash step, use a repeater pipette to fill each vessel with 1 ml of solvent and remove solvent by pushing it through the reactor vessel using the gas cover plate with nitrogen flow (Figure 3).
3. After washing, add 400 µl DMF to each vessel, move the entire U-Block Reactor assembly to a water bath sonicator, and sonicate for 30 min. To ensure thorough sonication without contamination, water levels in the sonicator should match the level of the fluid in the tubes. Depending on the size of the water bath sonicator, the reactor vessels may need to be transferred to a different rack.
In prior work, we have found that pushing solvents through vessels using this gas cover plate leads to more even washing than simply drawing solvents through using a vacuum manifold; however, this may depend on the precise setup used. To ensure even functionalization, during this and all subsequent steps, it is important to verify that the beads are uniformly distributed and don't clump. Check this frequently during sonication by observing the floating beads; if necessary, perform additional sonication steps to break up remaining clumps.

Part II: Functionalization of MRBLEs
Prior to peptide synthesis, the PEG-DA polymers that make up MRBLEs hydrogel beads must be chemically functionalized with free amine groups to allow direct coupling of Fmoc-protected amino acid building blocks. In this protocol, the outer shell of the MRBLEs is selectively functionalized with an acidresistant linker (which remains intact upon exposure to the TFA required for final side chain deprotection during Fmoc peptide synthesis) while the inner core is selectively functionalized with an acid-labile linker that is cleaved upon exposure to TFA (Kunys et al., 2012). In this manner, peptides from the inner core can be eluted during the final steps of peptide synthesis for direct analysis via mass spectrometry while peptides coupled to the outer shell remain attached for downstream binding assays. This procedure   Note: This step ensures that beads are swelled with water prior to performing the reaction in non-aqueous solvent.
3. Drain water carefully and make sure all the water around the beads is gone. There should still be water inside the beads for the next reaction and it is therefore important not to drain the beads for too long to prevent the inside of the beads from drying out.   If any air is found, wash lines to eliminate air bubbles. It will also show how much DMF is necessary for all the washing steps as well as the amount of 40% 4-Methylpiperidine. We generally make all the solutions in 10% excess to make sure there is enough (e.g., to account for evaporation during prolonged peptide synthesis). Prepare amino acids in 50 ml tubes. Fill HCTU and NMM in special glass peptide synthesizer bottles according to manufacturer's instructions. Fill DMF and 40% 4-Methylpiperidine glass bottles connected to peptide synthesizer.
ii. Start synthesizer. Synthesizing 48 peptides with 15 amino acids each takes approximately 24 to 48 h on the Syro II synthesizer, depending on coupling times and complexity of your library (e.g., whether a library contains systematic single amino acid changes of a particular peptide sequence vs. a collection of peptides with completely different sequences).

Part IV: Peptide loading density testing and characterization
Accurate multiplexed measurement of protein-peptide binding affinities requires that all reagents be present at approximately the same concentrations. Given that peptides are displayed on beads in the MRBLE-pep assay, this means that beads from each code are all approximately the same size, that equal numbers of beads are added to each assay, and that the surface density of displayed peptides is approximately constant. While microfluidically-produced MRBLE beads are significantly more monodisperse than most commercially available bead-based resins, care must be taken to ensure that each peptide sequence is displayed at approximately the same density. To directly visualize synthesized peptide density (and estimate peptide loading per bead), we biotinylate peptides to allow visualization using fluorescently labeled streptavidin via the following steps: (1) biotinylate the peptides after synthesis, (2) detect the biotinylated peptides using fluorescently labeled streptavidin and (3) decode the beads and measure fluorescence intensities ( Figure 5). Importantly, this step can be performed at any point in the synthesis protocol to allow troubleshooting of individual steps (e.g., after initial MRBLE functionalization or after particularly challenging coupling steps). In all cases, comparing fluorescence signals after a given protocol with appropriate negative controls can test that streptavidin binding is specific.
A. Biotinylation of MRBLE-bound peptides. 4. Take out about 10% of beads from each code and pool them in a new reaction vessel (e.g., draw 40 µl out of 400 µl after vigorous pipetting).

Part V: Peptide side chain deprotection and inner core peptide elution for sequence validation
A crucial advantage of bead-bound peptide libraries is that peptides can be eluted from beads and analyzed via mass spectrometry to ensure that any peptides showing decreased protein binding reflect a truly disrupted interaction rather than failure to synthesize the correct peptide. In this step, we describe elution of peptides from the inner core (by exposing this acid-labile linker to strong acid while removing side chain protecting groups during the final peptide synthesis step) followed by sequence analysis via MALDI mass spectrometry.  B. Transfer beads to 96 deep well plates or combine for downstream analysis. In most cases, we suggest leaving beads in separate vessels when possible to allow synthesis and substitution of individual additional peptides as required.

Part VI: Mass spectrometry to check quality of synthesized peptides
Peptide synthesis can be challenging and factors such as peptide length, amino acid repeats and choice of coupling reagents influence the success. In addition, quality control of synthesized peptides is especially important when peptides are synthesized directly on beads subsequently used for binding experiments since chromatographic purification steps are not possible. Here, we describe analysis of eluted peptides via mass spectrometry. We use MALDI-TOF for peptide analysis, as it has a high sensitivity and generates mostly single charged ionized peptides species that make downstream identification simple (Nadler et al., 2017). However, other mass spectrometry-based methods can be used as well.   However, several parameters must be considered carefully to ensure a successful binding experiment.
1. The number of beads added per code must be held constant within and between experiments.
Empirically, we find that ~100 beads per code ensures that the concentration of available peptide is sufficiently low relative to the concentration of available protein while simultaneously reducing variability due to stochastic Poisson error in the number of beads present in each volume. In addition, having 100 replicates of each measurement allows robust detection of small but statistically significant differences in binding between peptides via signal averaging.
2. Accurate quantification of binding requires that bound protein be detected via imaging. In past MRBLE-pep experiments, we detected binding of epitope-tagged proteins using an antibody . Here, we describe quantitation of binding for proteins that are directly tagged with a fluorescent indicator, e.g., YFP. In general, any tagging strategy can be used as long as appropriate negative controls are included to eliminate spurious fluorescence that derives from nonspecific interactions. For binding assays using fluorescent protein tags, an appropriate negative 20 www.bio-protocol.org/e3669 control should include the fluorescent protein alone. When using fluorescently labeled antibodies, binding assays should: (1) incubate the protein of interest 1:1 with antibody prior to incubation with beads to reduce direct binding of free labeled antibody to beads, and (2) directly measure binding of the labeled antibody alone to peptide-bead libraries.
3. The amount of protein added should include a concentration range that is 10-fold higher and lower than the expected interaction Kd whenever possible. We typically use a range of protein concentrations spanning from ~30 nM to 2 µM. However, this may need to be adjusted depending on expected affinities or the presence of protein aggregation at high concentrations. 1. For each experiment, use ~100 beads per code. In the protocol described here, this corresponds to ~1% of the beads from each code (e.g., if beads of 1 code are resuspended in 1 ml of buffer, draw up 10 µl of beads after extensive mixing). In typical assays, this means measurements take place on ~4,800 beads for 48 codes in a total volume of 20 µl.
2. Spin down beads, remove PBS-T and block to prevent excess nonspecific binding by incubating with 100 µl 5% BSA in PBS-T for 1 h rotating at room temperature at 20 rpm. 3. Use microManager and multidimensional acquisition to take multiple images to cover entire area with beads at all Lanthanide channels and exposure times, brightfield and assay channel. Here, we used an YFP filter cube (Semrock, Rochester) to detect YFP-tagged Calcineurin and a Cy5 filter cube (Semrock, Rochester) to detect Dylight 650 tagged Streptavidin. 4. Images were analyzed using the Python code available for download on Github . The images were loaded into the software analysis package and then processed to decode beads and obtain fluorescence intensities for each bead and each code. The Excel table was saved and used to further process and plot the data with Python ( Figure 9). 5. We used Python to plot the mean background subtracted fluorescence intensities for indicated peptides as well as fit the Langmuir isotherm ( Figure 10). 22 www.bio-protocol.org/e3669   values. The left plot shows curves for a known calcineurin substrate (PRIIIT, Kd = 1.67 ± 0.86 µM calculated as the mean ± standard deviation of the returned values from each code using a global fit); the right plot shows curves for a negative control peptide.

Notes
We use 48 x 2 ml reaction vessels and a U-Block reactor equipment, which are directly compatible with the Syro II peptide synthesizer used for high-throughput peptide synthesis. However, this protocol can be adapted to any reaction vessels and mounting plates. The reaction vessels are simple columns with a filter. The U-Block holds the reaction vessel and the bottom of each vessel holder is connected to tubing which allows washing of the vessels. The vessels can be washed by either applying a vacuum or pushing the liquid through the vessel by using the inert gas plate.