A Mass Spectrometry-Based Proteomics Approach for Global and High-Confidence Protein R-Methylation Analysis

1. Cell culturing and protein extraction (time: 3 – 4 weeks required)

1. Grow HeLa cells in parallel in media supplied with either Light (L) or Heavy (H) Methionine, respectively (see Table 1 for media composition). Upon at least eight cell divisions, collect an aliquot of cells from each SILAC channel and perform the incorporation test.
   NOTE: To check for the incorporation efficiency, test by LC-MS/MS analysis that the percentage of heavy Methionine (Met-4) in the Heavy channel is as near as possible to 100%. Analyze an aliquot of heavy-labeled cells by LC-MS/MS (for settings see Table 2), then process the MS data with MaxQuant using the parameters indicated in Table 3. To check for the Met-4 incorporation an in-house developed script is available at https://bitbucket.org/EMassi/hmseeker/src/master/.
2. Consider heavy Methionine incorporation as complete when it reaches >97%. When each channel reaches a total number of about 60 x 10⁶ of cells (corresponding to about 40 dishes of 15 cm each at 85% confluency for HeLa cells, with variations depending on the cell type) harvest them. Carefully count them, mix in 1:1 proportion and pellet by centrifugation at 335 x g for 5 min at 4 °C.
   NOTE: To assess the proper 1:1 L/H mixing, keep an aliquot and run it on a slice of a gel which is known to contain a high abundance and heavily R-methylated protein (e.g., fibrillarin). If labeling has been successful and 1:1 mixing has been achieved, there should be a 1:1 ratio of light and heavy versions of the R-methylated peptides present in the sample. Alternatively, keep an aliquot of mixed sample to be analyzed by LC-MS/MS, then process the MS data with MaxQuant using the parameters indicated in Table 3 and plot the distribution of Log2 H/L ratio as depicted in Figure 2C. The protocol can be stopped here by snap-freezing the pellet and storing it at -80 °C.
3. Re-suspend the cell pellet in four volumes of Lysis Buffer (see Table 1 for Lysis Buffer composition) with respect to the cell pellet volume. For instance, use 6 mL of Lysis Buffer for a pellet from 120 x 10⁶ Hela cells (60 x 10⁶ Light + 60 x 10⁶ Heavy) corresponding to 1.5 mL volume.
   NOTE: Protein extraction must be performed at room temperature (RT) because Lysis Buffer contains the chaotropic agent 9 M Urea that precipitates at ice temperature; therefore, the addition of a broad spectrum of Serine and Cysteine protease inhibitors is important, as well as phosphatases inhibitor, to simultaneously protect proteins against proteolytic degradation and dephosphorylation, cocktail of protease and phosphatase inhibitors are commercially available as small tablets, see Table 1 and Table of Materials.
4. Sonicate the sample with a microtip cell disruptor sonicator for at least five cycles of 15 s ON and 30 s OFF, to ensure efficient breakage of cell membranes and DNA release and shearing. Check the viscosity of the extract by pipetting the solution up and down. If it is too viscous due to incomplete DNA shearing and membrane solubilization, repeat the sonication cycles.
   NOTE: Ensure that the sample does not over-heat during sonication, because high temperature can damage proteins. However, it is not possible to put the sample on ice between sonication cycles, because of the presence of 9M Urea; hence, it is advisable to pause for 60 s OFF between different sonication cycles. Moreover, avoid the formation of air bubbles during sonication because they reduce the sonication efficacy.
5. Centrifuge the extract at 3,000 x g for 10 min at RT to pellet the debris and transfer the supernatant in a new 15 mL tube.
6. Measure the protein content of the extract with a colorimetric assay, such as Bradford or bicinchoninic acid (BCA)^14,15. An optimal starting amount of protein extract for this protocol is between 20-30 mg.
   NOTE: Lysis buffers containing high concentration urea are compatible both with Bradford and BCA quantification assay; other types of Lysis buffer, such as those including high concentration of sodium dodecyl sulphate (SDS), are not compatible with Bradford.

2. Lysate digestion (indicative time required 2 hours)

1. Perform reduction of thiol group (-SH) of proteins using a stock solution of dithiothreitol (DTT) dissolved in ultrapure water at a final concentration of 4.5 mM and let the reaction go for 30 min at 55 °C.
   NOTE: It is possible to prepare 1 M stock DTT solution and store it at -20 °C for up to 1 month, thawing just the aliquots needed for each experiment. Alternatively, sulfhydryl reductant tris-(2-carboxyethyl)-phosphine (TCEP) can be used to perform reduction of -SH groups; especially for long-term storage of proteins, TCEP is significantly more stable than DTT without metal chelates such as EGTA in the buffer, whereas DTT is more stable if metal chelates are present¹⁶.
2. Perform alkylation of thiol group (-SH) of proteins by adding iodoacetamide (IAA) at a concentration of 10 mM and incubate for 15 min at RT in dark. Perform the incubation of extracted proteins with IAA solution in dark because IAA is photosensitive.
   NOTE: The IAA stock solution at 100 mM should be prepared fresh before each experiment. Alternatively, chloroacetamide could be used to perform alkylation of -SH groups, especially if the goal of the experiment is to analyze cross-talk between methylation and ubiquitination because IAA-induced artefact mimics ubiquitination¹⁷.
3. Before proceeding with the protein digestion step, save an aliquot of protein extract (1/1,000 of starting undigested lysate) for subsequent analysis on SDS-PAGE Coomassie-stained gel and comparison with a corresponding amount of sample upon digestion; this test serves to verify the proteolysis efficiency (see point 4).
4. Dilute the remaining protein extract with four volumes of 20 mM HEPES pH 8.0, to reach a final urea concentration of 2 M (which is the concentration compatible with the enzymatic activity of proteases). Split the sample into two parts: in the first add Sequencing Grade Modified Trypsin and in the second add LysargiNase protease (see Table of Materials) at 1:100 (w/w) proportion relative to the mg of starting material. Leave overnight at 37 °C in a thermomixer at 600 rpm, to allow enzymatic digestion.
   ​NOTE: Trypsin, the most common digestion enzyme in proteomics, cleaves at the C-terminus of R and Lysine (K), generating peptides with a charge distribution that results in fragmentation spectra dominated by y-type ions upon collision-induced dissociation (CID). LysargiNase cleaves at the N-terminus of R and K, therefore, mirroring the Trypsin cleavage specificity and generating peptides that release mainly b-type ions upon CID fragmentation. This combined analysis leads to much increased peptide sequence coverage and in higher confidence in site-specific identification of R-methylations¹⁸.

3. Peptide purification (indicative time required 1 hour)

1. Keep an aliquot of digested peptides from both reactions, collecting the same volumes as in point 2.3 for the comparison on SDS-PAGE Coomassie-stained gel to assess protease digestion efficiency (see point 4).
2. Stop the digestion by acidifying the samples with the addition of trifluoroacetic acid (TFA) to a final concentration of 5%. Mix well and measure the samples pH with a litmus paper (pH should be around 3). Vortex briefly and spin down the acidified samples before transferring them into new 15 mL tubes.
3. Clean up the samples through two C18 vac cartridge (sorbent weight 1 g, see Table of Materials), one for the sample digested with Trypsin and the other for the sample digested with LysargiNase. Prepare Solvent A, Solvent B, and Wash Buffer (see Table 1 for buffer composition).
4. Using glass pipettes, rehydrate each cartridge with 6 mL of ACN 100% for 3 times. After that, equilibrate each cartridge sequentially with 3-9-18 mL of Solvent A. Load the samples (the resins should become yellow). Wash again sequentially with 3-9-18 mL of Solvent A and then add 6 mL of Wash buffer. Transfer each column into clean 15 mL tubes and elute the samples with 7 mL of Solvent B. Repeat the elution step with 7 mL of Solvent B, for a final volume of 14 mL.
   NOTE: Perform all these steps by letting the buffers and solution pass through the columns by gravity. To favor the flow of the buffers through the column, push each solution slowly with a syringe, to mimic vacuum.
5. Save 50 µL of eluted peptides, 50 µL of flow-through (FT), 50 µL of the wash with Solvent A, and 50 µL of the last wash for the subsequent peptide assessment by SDS-PAGE (see point 4).

4. Coomassie-stained SDS-PAGE gel (indicative time required 2 hour)

1. Run the collected aliquots on a 17.5% SDS-PAGE gel and stain with Instant-Blu Coomassie staining (see Table of Materials). The expected result is represented in Figure 2A.

5. Peptide lyophilization (indicative time 2 days)

1. Cover the 15 mL tubes containing the eluted peptides with paraffin film, which is then punched with a 20 G needle to create 3-5 holes. Put the tubes in dry ice for at least 30 min, until the samples are completely frozen.
2. Lyophilize the frozen fractions for 48 h, a time interval typically sufficient to ensure a complete lyophilization of the samples, even if some variability may occur, due to the freeze dryer performance.
   ​NOTE: The experiment can be paused here, storing the lyophilized samples at -80 °C.

6. Off-line HpH-RP chromatographic fractionation of peptides (indicative time 4 days)

1. To fractionate the peptides into 60 fractions, use HpH-RP liquid chromatography, using HPLC system equipped by C12-RP HPLC column (250 x 4.6 mm, 4 µm Proteo 90A).
2. Before the run, prepare fresh Buffer A and Buffer B (the composition of the Buffers is described in Table 1).
3. Filter all solution with 0.22 µm filter and degas them in a sonicator bath for at least 30 min.
4. Dissolve the lyophilized peptides in 1 mL of Buffer A. Filter the peptides through a polytetrafluoroethylene (PTFE) 0.45 µm filter, using a syringe.
5. Set the fractionation rate at 1 mL/min flow and collect 1 mL of fractions, using the following chromatographic gradient: 5% B to 30% B in 60 min; 30% B to 60% in 2 min; 70% B for 3 min.
6. Set the HPLC so that, at this point, fraction collection is halted, and the gradient held at 70% Buffer B for 5 min before an extensive wash of the column with a quick gradient up to 100% Buffer B, followed by a final wash (100% Buffer B for 10 min).
   ​NOTE: At the end of each chromatographic run, always equilibrate the column with 100% Buffer A for 20 min.
7. Fractionate both samples separately digested with Trypsin and LysargiNase by Off-Line HpH RP chromatographic gradients, as described at point 6.5.
8. For each chromatographic gradient, collect all the fractions into a deep 96 well plate.
9. Pool the fractions collected before the gradient into one single fraction named PRE. Concatenate the 60 fractions from the HpH-RP liquid chromatographic (LC) gradient by pooling them in a non-contiguous way into 14 final fractions. To obtain such non-contiguous concatenation, pool the HpH-RP fractions according to the following scheme.
   1. Fraction 1 (final volume 5 mL): Pool 1-15-29-43-57
   2. Fraction 2 (final volume 5 mL): Pool 2-16-30-44-58
   3. Fraction 3 (final volume 5 mL): Pool 3-17-31-45-59
   4. Fraction 4 (final volume 5 mL): Pool 4-18-32-46-60
   5. Fraction 5 (final volume 4 mL): Pool 5-19-33-47
   6. Fraction 6 (final volume 4 mL): Pool 6-20-34-48
   7. Fraction 7 (final volume 4 mL): Pool 7-21-35-49
   8. Fraction 8 (final volume 4 mL): Pool 8-22-36-50
   9. Fraction 9 (final volume 4 mL): Pool 9-23-37-51
   10. Fraction 10 (final volume 4 mL): Pool 10-24-38-52
   11. Fraction 11 (final volume 4 mL): Pool 11-25-39-53
   12. Fraction 12 (final volume 4 mL): Pool 12-26-40-54
   13. Fraction 13 (final volume 4 mL): Pool 13-27-41-55
   14. Fraction 14 (final volume 4 mL): Pool 14-28-42-56
       NOTE: The non-contiguous concatenation consists in combining early-, mid-, and late-eluting fractions, which allows increasing the heterogeneity in peptide composition within the pooled fractions. Consequently, the peptide mixture of each pooled fraction is efficiently separated, with limited co-elution, in the subsequent nano-flow low pH-RP-LC chromatography directly coupled to the mass spectrometer.
10. Pool the fractions collected after the gradient into a unique fraction, named POST.
    NOTE: By including the fractions PRE and POST gradient, a total of 16 fractions are obtained, in 15 mL tubes (see Figure 3A).
11. Cover the 15 mL tubes with paraffin film and punch it with a 20 G needle to generate 3-5 holes. Freeze them by incubating the centrifuge tubes in dry ice until each fraction is completely frozen.
12. Lyophilize the fractions for about 48 h. Ensure that each sample is completely dried before stopping the freeze-dryer.
    NOTE: The experiment can be paused here, storing the lyophilized samples at -80 °C.

7. R-methylated peptide immuno-affinity enrichment (indicative time 2 days)

1. Perform the sequential immuno-affinity enrichment of modified peptide with anti-pan-R-methylation antibodies in parallel, but separately for the two samples from Trypsin and LysargiNase digestions, respectively. The Immuno-Affinity Purification (IAP) Buffer is provided by the company purchasing the anti-pan-R-methyl antibodies for modified peptide affinity enrichment (details are in Table Material and Reagents). The IAP buffer is concentrated 10x and should be diluted 10 times before use.
   NOTE: The IAP Buffer 1x can be stored at -20 °C to up to 1 year.
2. Centrifuge the lyophilized peptides at 2,000 x g for 5 min at RT to spin down the peptides to the bottom of the 15 mL tube. Re-suspend the lyophilized peptides with 250 µL of 1x IAP Buffer per 15 mL tube and transfer in a 1.5 mL low-binding tube. Check using a litmus paper whether the pH is >6.
3. Keep a small aliquot (about 5% of the volume) of each fraction as input for the subsequent MS analysis.
4. Split each fraction in two aliquots, in order to perform the immuno-enrichment of asymmetrically-di-methylated (ADMA) and symmetrically-di-methylated (SDMA) peptides in parallel.
5. Use three vials of the selected anti-pan-R-methylated antibodies conjugated to protein A agarose beads per 10 mg of the initial protein extract.
6. Prepare the correct amount of antibody conjugated to agarose beads by centrifuging each vial at 2,000 x g for 30 s and removing the buffer from the beads. Wash the beads three times with 1 mL of 1x PBS always by centrifuging them at 2,000 x g for 30 s.
7. After the last wash, re-suspend the beads in 40 µL 1x PBS for each vial; pool them and finally divide them equally into 16 fractions (so that 2.5 µL of antibody-beads is added to each fraction).
8. Add 250 µL of 1x IAP Buffer to each tube, mix by inverting and let it incubate on a rotating wheel for 2 h at 4 °C.
   NOTE: Mix the samples by inverting the 1.5 mL tubes rather than by pipetting them with microtips, which could damage the beads or result in losing them.
9. Upon 2 h incubation, centrifuge the 1.5 mL tubes containing peptides and pan-R-methyl-antibody-conjugated beads at 2,000 x g for 30 s to pellet the beads; transfer the FT from each fraction into clean 1.5 mL low-binding tubes.
10. Add the beads conjugated to antibodies against R-mono-methylation (MMA) to the FTs and repeat the steps 7.7 to 7.9.
11. During the incubation of the peptide samples with the MMA-beads, wash twice the fractions which were previously immuno-precipitated with anti-ADMA and SDMA with 250 µL IAP Buffer (inverting and not pipetting), and discard the supernatant at each wash.
12. Repeat the wash with LC-MS grade H₂O thrice.
13. Elute the affinity-enriched symmetrically and asymmetrically R-di-methylated peptides from the agarose beads by adding 50 µL of 0.15% TFA to each tube (strong acid conditions, in fact, denature the epitope leading to the release of the antigens from the antibodies). Leave this solution 10 min at RT, inverting the tubes every 2-3 min.
14. Transfer the first elution into clean 1.5 mL low-binding tubes and repeat the elution with 50 µL 0.15% TFA; pool the 2 fractions in one tube.
15. Repeat steps from 7.11 to 7.14 for the R-mono-methylated peptides that were incubated with the anti-MMA antibody-beads.

8. Desalting and concentration of affinity-enriched methyl-peptides by C18 microcolumns (indicative time required 30 minutes)

1. Equilibrate with methanol the C18-RP microcolumns made with 3M Solid Phase extraction cartridges for peptide desalting and concentration prior to MS analysis¹⁹.
2. Load the samples (corresponding to the separate immuno-affinity enriched fractions and input fractions) on the C18 microcolumns in two steps (50 µL + 50 µL on each C18 microcolumn) by centrifuging at 600 x g for 6 min.
3. Wash the microcolumns with 55 µL Buffer A (see Table 1 for buffer composition), always by centrifugation at around 900 x g for 5 min.
   NOTE: The experiment can be paused here, leaving the C18-RP microcolumns at 4 °C, where they can be stored up to 2 weeks.

9. Second enzymatic digestion (indicative time required 3 hours)

1. Wash the C18-RP microcolumns with 55 µL of Buffer A (see Table 1 for buffer composition) for two times, by centrifugation at around 850 x g for 5 min.
2. Elute the peptides twice with 20 µL of Buffer B (see Table 1 for buffer composition) and pool the two fractions.
3. Dry the eluted peptides in a vacuum concentrator (see Table Material and Reagents for details). Meanwhile, prepare the digestion solution that consists of 50 mM ammonium bicarbonate that is diluted from a freshly made 1 M stock solution (see Table 1).
4. Add Trypsin or LysargiNase to the respective samples, to a final concentration of 25 ng/µL. Incubate each sample at 37 °C for 2 h.
5. Add 1 µL of 5% TFA to stop digestion; vortex and spin down the samples.
   ​NOTE: Enzymatic cleavage by Trypsin can be inhibited at the C-terminus of methylated R and K, causing missed cleavages that increase peptide length and charge, which in turn produce complex and incomplete fragmentation spectra hindering peptide identification and site-specific attribution of methylation sites. It has been shown that a second enzymatic digestion may reduce the frequency of such missed cleavages, with improved sequence coverage and site-attribution²⁰.

10. Desalting peptides (indicative time required 30 minutes)

1. Load the acidified peptide solutions to new C18-RP microcolumns that have been previously equilibrated following the same steps described in point 8.
2. The peptide loaded on the C18-RP microcolumns can be stored at 4 °C until elution for LC-MS/MS analysis.

11. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis (indicative time 5 days)

1. Elute the peptides from the C18-RP microcolumns by passing 10µL of Buffer B, centrifuging at 615 x g for 5min at RT. Repeat this step twice and combine the eluates.
2. Reduce the volume of the eluates in a vacuum concentrator until they are almost dry, avoiding over drying.
3. Re-suspend the peptides in 10 µL of Buffer A for LC-MS/MS analysis.
4. Analyze each fraction of R-methyl-peptides by liquid chromatography-tandem mass spectrometry (LC-MS/MS) in a high-resolution Mass Spectrometer (see Table of Materials), coupled to a nano-flow ultra-high-performance liquid chromatography (UHPLC) system. Set the instrument parameters as described in Table 2.
5. Load 2 µL of each sample on a nano-analytical column (easy spray column 75 µm inner diameter, 25 cm length), packed with C18-RP resin (2 µm particle size).
6. Samples are passed through the C18 RP nano-column at a flow rate of 300 nL/min, with the following linear gradient: 3%-30% B for 89 min, 30%-60% B for 5 min, 60%-95% B for 1 min, and 95% B for 5 min.
7. The mass spectrometer operates in data-dependent acquisition (DDA) mode to automatically switch between full scan MS and MS/MS acquisition. Set the survey full scan MS to be analyzed in the spectrometer detector with resolution R = 70,000. The fifteen most intense peptide ions are sequentially isolated to a target value of 3 x 10⁶ and fragmented by relative collision energy of 28%. Set the maximum allowed ion accumulation times to 20 ms for full scans and 50 ms for MS/MS and fix the target value for MSMS to 1 x 10⁶. The dynamic exclusion time is set to 20 s.

12. Running MaxQuant and hmSEEKER data analysis

1. Upon completion of the LC-MS/MS runs, import the MS raw data into a peptide search engine to identify the methyl-peptides by probability-based approach against the reference database. In this protocol, MaxQuant version 1.6.2.10 was used for our analysis. MaxQuant requires a minimum of 2 GB RAM to run, as well as enough disk space to store all the raw data and all the output files.
   NOTE: Refer to the official documentation at https://www.maxquant.org for all the details about the installation and the hardware and software requirements.
2. Duplicate each raw data file. Rename the originals by appending "_light" to their name, then rename the copies by appending "_heavy".
   NOTE: hmSEEKER, the script for downstream analysis, is case sensitive.
3. Launch MaxQuant/Andromeda search for peptide identification with the settings indicated in Table 3. Of the several output data produced by MaxQuant, only the allPeptides.txt and msms.txt files (located in the combined/txt subfolder) are required for the post processing step.
4. The post processing of MaxQuant output data is carried out by the algorithm hmSEEKER. Download hmSEEKER from: https://bitbucket.org/EMassi/hmseeker/src/master/. The script is available as a Jupyter notebook written in Python 3.7 and comes with a sample dataset for testing purposes. For new users, it is advisable to download and install the Anaconda platform (https://www.anaconda.com/products/individual). The latest release includes by default Python 3.8, Jupyter and all the packages that are required to run hmSEEKER (e.g., Scikit-learn 0.23.1).
5. Create a folder and store the files allPeptides.txt and msms.txt from MaxQuant output into it.
6. Launch Jupyter (from the command line or from the Anaconda navigator).
7. Navigate to the hmSEEKER folder and open hmSEEKER.ipynb.
8. In the Input Parameters section of the notebook, indicate the paths to the FASTA database and to the folder(s) containing the MaxQuant text files.
9. Run the code inside each cell by selecting the cell and clicking on the Play button on top of the Jupyter interface.
10. The script produces a comma-separated output file for each dataset that was analyzed, plus a combined file. The final doublets list can be found in the file named "[date]-[time]-combined_hmSILAC_doublets_HxL_summary.csv"
    (Table 4 includes a brief description of the columns in the output table).