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. Author manuscript; available in PMC: 2023 Jan 1.
Published in final edited form as: Methods Mol Biol. 2022;2420:21–37. doi: 10.1007/978-1-0716-1936-0_3

Proteomic Profiling of Cerebrospinal Fluid by 16-Plex TMT-Based Mass Spectrometry

Kaushik Kumar Dey, Huan Sun, Zhen Wang, Mingming Niu, Hong Wang, Yun Jiao, Xiaojun Sun, Yuxin Li, Junmin Peng
PMCID: PMC8890903  NIHMSID: NIHMS1771189  PMID: 34905163

Abstract

Mass spectrometry (MS) has become a mainstream platform for comprehensive profiling of proteome, especially with the improvement of multiplexed tandem mass tag labeling coupled with two-dimensional liquid chromatography and tandem mass spectrometry (TMT-LC/LC-MS/MS). Recently, we have established a robust method for direct profiling of undepleted cerebrospinal fluid (CSF) proteome with the 16-plex TMTpro method, in which we optimized parameters in experimental steps of sample preparation, TMT labeling, LC/LC fractionation, tandem mass spectrometry, and computational data processing. The extensive LC fractionation not only enhances proteome coverage of the CSF but also alleviates ratio distortion of TMT quantification. The crucial quality control steps and improvements specific for the TMT16 analysis are highlighted. More than 3000 proteins can be quantified in a single experiment from 16 different CSF samples. This multiplexed method offers a powerful tool for profiling a variety of complex biofluids samples such as CSF, serum/plasma, and other clinical specimens.

Keywords: Cerebrospinal fluid, Serum, Plasma, Proteomics, Proteome, Clinical proteomics, Liquid chromatography, Mass spectrometry, Isobaric labeling, Tandem mass tag

1. Introduction

Cerebrospinal fluid (CSF) is a body fluid circling in the brain and spinal cord, which is reflective of the underlying pathological state of the central nervous system (CNS). The CSF circulates within the ventricular system of the brain and spinal cord, consisting of low concentration of proteins derived from blood, central nervous system, and peripheral tissue [1–4]. The CSF shows a unique advantage over plasma and other fluid sources to reveal biochemical and pathological changes of the brain [5, 6]. However, comprehensive profiling of CSF proteome is often difficult due to a large dynamic range of protein concentrations, as exemplified by a small subset of highly abundant proteins [7, 8]. To reduce protein dynamic range, antibody-based depletion of the most abundant proteins is often utilized to enhance the detection of low abundance proteins [9–11], but the depletion step introduces additional experimental variations [11]. Alternatively, the challenge of dynamic range can be alleviated in the established proteomics platform that combines the latest multiplexed tandem mass tag (TMT) method, extensive fractionation by two-dimensional liquid chromatography (LC/LC), and high-resolution tandem mass spectrometry (MS/MS) method [12]. Recently, our group performed an ultra-deep discovery study by 11-plex TMT-LC/LC-MS/MS to bypass the depletion step, detecting 5941 and 4826 proteins in human CSF and serum, respectively [12–14]. Potential biomarker proteins were further analyzed in validation experiments by a targeted LC-MS3 method (e.g., TOMAHAQ: Triggered by Offset, Multiplexed, Accurate mass, High resolution, and Absolute Quantitation) [12, 15]. With substantial technical improvements, TMT-based quantitative mass spectrometry has become a mature platform for high throughput and deep profiling of proteome and protein modifications [13, 16–18]. In this platform, peptide fractionation can be achieved by the integration of basic pH reverse phase LC and acidic pH reverse phase LC, which improves peptide separation power and alleviates TMT reporter ion ratio distortion [19, 20] during mass spectrometric data acquisition and analysis [12, 21–23]. More recently, TMTpro (termed TMT16 hereafter) reagents further increase the multiplex capacity [24–26]. The TMT16 reagents contain a proline-based reporter group, a mass balance group, and an amine reactive group (Fig. 1a, b). To enable the combination of stable C13 and N15 isotopes to achieve 16-plexing, the reporter and mass balance groups of TMT16 are significantly modified from the original TMT6–11 reagents. The additional channels in the TMT16 also increase the total amount of the starting material when pooling TMT-labeled samples, which is beneficial to the analysis of limited samples [27]. Our previous published articles have provided an in-depth overview of the TMT10–11 platform [12, 14, 21, 28]. In this protocol, we describe a comprehensive pipeline for profiling human CSF proteome (Fig. 2). CSF proteins are digested into peptides, labeled with TMT16 reagents, pooled in an equal ratio, and analyzed by LC/LC-MS/MS. Finally, the JUMP software suit is used for data analysis [19, 29, 30] (see Note 1). This protocol can be regularly used to identify and quantify ~3000 proteins from CSF proteome with high precision. Thus, this newly established method is a general application for deep analysis of CSF and other biofluids.

Fig. 1.

Fig. 1

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Structure of the 16-plex TMT reagent. (a) Structure of the 16-plex TMT reagent, labeling process, mass shift after labeling, and the mass of the reporter ion are shown. (b) Heavy isotope–labeled structures of the reporter ions of TMT16 reagents

Fig. 2.

Fig. 2

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Experimental scheme of 16-plex TMT-LC/LC-MS/MS for profiling CSF whole proteome

2. Materials

2.1. CSF Protein Extraction and In-Solution Digestion

  1. Lysis buffer: 50 mM [4-(2-hydroxyethyl)-1-piperazineethane-sulfonic acid] (HEPES), pH 8.5, 8 M urea (see Note 2), 0.5% sodium deoxycholate, 1 × PhosSTOP phosphatase inhibitor cocktail.

  2. 20 mg/mL bovine serum albumin (BSA).

  3. 10% sodium dodecyl sulfate (SDS) polyacrylamide gel.

  4. GelCode Blue Stain.

  5. BCA Protein Assay Kit (Thermo Fisher Scientific).

  6. 1 M dithiothreitol (DTT) in 50 mM HEPES.

  7. 1 M iodoacetamide (IAA) in 50 mM HEPES (see Note 3).

  8. Acetonitrile (ACN, HPLC grade).

  9. Lys-C (FUJIFILM Wako).

  10. Trypsin (Promega, see Note 4).

  11. Trifluoracetic acid (TFA).

2.2. Peptide Desalting

  1. C18 ZipTips (Millipore).

  2. C18 Desalting column (Ultra MicroSpin Columns): loading capacity of ~120 μg peptides (Harvard Apparatus 74–7243 ~ 20 μL of bed volumes) (see Note 5).

  3. Vacuum manifold.

  4. Methanol (HPLC grade).

  5. Equilibration and wash buffer: 0.1% TFA in water.

  6. Elution buffer: 60% ACN and 0.1% TFA.

  7. Savant SpeedVac concentrator (Thermo Scientific).

2.3. TMT Labeling of Peptides and Desalting

  1. 50 mM HEPES, pH 8.5.

  2. 16-plex TMTpro Isobaric Mass Tagging Kit (Thermo Fisher Scientific).

  3. Anhydrous ACN.

  4. C18 ZipTips (Millipore).

  5. Desalting column (cartridge): 100 mg Sep-Pak C18 desalting cartridge (Waters, ~1.1 mL of bed volumes) (see Note 5).

  6. Methanol (HPLC grade).

  7. Equilibration buffer: 0.1% TFA in water.

  8. Washing buffer: 5% ACN and 0.1% TFA.

  9. Elution buffer: 60% ACN and 0.1% TFA.

  10. TMT quenching solution: 5% hydroxylamine.

2.4. Offline Basic pH Reverse Phase Liquid Chromatography (RPLC) Fractionation

  1. Waters XBridge C18 column, 4.6 mm × 25 cm, 3.5 μm particle size (see Note 6).

  2. Buffer A: 10 mM ammonium formate, adjust pH to 8.0 by 28% ammonium hydroxide.

  3. Buffer B: 10 mM ammonium formate, 90% ACN, adjust pH to 8.0 by 28% ammonium hydroxide.

  4. Washing solution: 25% isopropanol, 25% methanol, 25% ACN, and 25% water.

  5. High-performance liquid chromatography (HPLC) system (e.g., Agilent 1220 infinity LC system).

  6. Fraction collector (e.g., Gilson FC 203B)

2.5. Acidic pH RPLC-MS/MS Analysis

  1. Empty columns (New Objective, 75 μm I.D. × 15 cm, 15 μm tip orifice).

  2. C18 resin (Dr. Maisch GmbH Germany, 1.9 μm particle size) (see Note 6).

  3. Butterfly portfolio column heater.

  4. HPLC system (e.g., Waters ACQUITY UPLC or Dionex Ultimate 3000 RSLC nano system).

  5. Tandem MS instrument (e.g., Thermo Q Exactive HF or Fusion).

  6. Sample loading solution: 5% formic acid in water.

  7. Sample vials with 0.2 mL bottom-springed polypropylene inserts.

  8. Buffer A: 3% dimethyl sulfoxide and 0.2% formic acid (see Note 7).

  9. Buffer B: 67% ACN, 3% dimethyl sulfoxide, and 0.2% formic acid.

2.6. MS Data Analysis

  1. Database downloaded from UniProt website (https://www.uniprot.org/downloads).

  2. Proteomics software suite for data processing: JUMP [19, 29, 30] or Proteome Discover and Bio works (Thermo Scientific).

  3. A computer cluster for data processing.

3. Methods

3.1. Cerebrospinal Fluid

Human CSFs were provided by the brain bank supported by the brain and body donation program at Banner Sun Health Research Institute with well-established criteria for clinical and pathological diagnoses [31, 32]. All subjects consented to the study. All samples were frozen and stored in aliquots of polyethylene tubes at −80 °C until use. The CSF samples should be colorless. Blood contamination may be found by the red color of the samples.

3.2. Protein Extraction, Quality Control, and In-Solution Digestion

All the procedures are performed at room temperature (21 °C) unless otherwise specified, as high concentration of urea in the lysis buffer may crystalize at 4 °C.

3.2.1. Protein Extraction and Quality Control

To reduce the quantitative variations introduced during sample processing, it is critical to follow the experimental procedures and handle each sample consistently. The CSF proteins are denatured in fresh lysis buffer containing 50 mM HEPES, pH 8.5, 8 M urea, 0.5% sodium deoxycholate, and 1 × phosphatase inhibitor cocktail. The Bicinchoninic acid assay (BCA) or short SDS gel staining method [33] is used for the quantification of protein concentration. The protein lysates are stored at −80 °C in aliquots for further use. During sample lysis, it should be noted that urea itself also occupies volume (100 mg of urea occupies ~73 μL in solution) and the CSF volume should be considered. The protein concentration of CSF is ~0.1–0.5 mg/mL.

  1. Make aliquots for CSF samples. Small aliquots (~20 μL) are used for the analysis of protein concentration, the evaluation of protein quality, and positive control proteins by Western blotting. Large aliquots (~200 μL) are used for proteomics analysis. Freeze the aliquots immediately on dry ice and store at −80 °C.

  2. Protein concentration can be evaluated by running a short SDS gel with the standard BSA (e.g., BSA titrations of 0.1, 0.3, 1.0, and 3.0 μg) [33]. All protein samples (each of ~1.0 μg) are analyzed in a 10% SDS gel until the running dye runs ~4 mm into the gel. The gel is stained with GelCode Blue stain buffer for 1 h and further destained with deionized H2O until the gel background is clear. The gel is then scanned and quantified by the ImageJ program (NIH). Based on the working curve of BSA standard, the absolute protein amount in each sample is calculated. Alternatively, the BCA can also be used for protein quantification (see Note 8).

  3. Quality control: analyze ~1.0 μg of each sample from the small aliquot by a gradient SDS gel (e.g., 4–20%) until bromophenol blue dye reaches the bottom of the gel. Stain the gel with GelCode Blue. Evaluate protein quality and remove highly degraded samples (see Note 9).

  4. Prepare pre-cooled 1.5-mL Eppendorf tubes over dry ice, and CSF samples are transferred immediately for extraction and digestion.

  5. Add solid urea and concentrated stock solutions to make lysate with expected final concentration (50 mM HEPES, pH 8.5, 8 M urea, 0.5% sodium deoxycholate, and 1 × PhosSTOP, see Note 10). It is recommended to add phosphatase inhibitors into the lysis buffer if analyzing phosphoproteins.

  6. Tighten up the caps and vortex the samples for 30 s. Confirm that urea is fully dissolved.

3.2.2. In-Solution Protein Digestion, Peptide Reduction and Alkylation, Digestion Efficiency Test, and Peptide Desalting

Protein digestion is an essential step for shot-gun proteomics [34], and in-solution digestion is the simplest method in terms of sample handling and speed. Trypsin is the most frequently used protease to produce peptides in the preferred mass range, which generates information-rich, interpretable MS/MS spectra. Prior to trypsin digestion, Lys-C is regularly used under the harsh lysis condition (8 M urea) to digest fully denatured proteins. Then the samples are diluted to 2 M urea for trypsinization. The two-step digestion improves digestion efficiency. In addition, Cys residues are reduced by dithiothreitol (DTT) and alkylated by iodoacetamide (IAA).

  1. Add 10% ACN and perform the Lys-C digestion into the large aliquot of CSF lysate (at least 20 μg) at an enzyme-to-substrate (protein/Lys-C) ratio of 1:50 (w/w) and incubate at room temperature for 3 h to reduce protein disulfide bonds.

  2. Dilute samples to 2 M urea with 50 mM HEPES (pH 8.5), and add freshly prepared 1 M DTT to final 1 mM.

  3. Add trypsin into each diluted sample at an enzyme-to-substrate (protein/trypsin) ratio of 1:20 (w/w) and digest at 21 °C for 3 h or overnight.

  4. Add DTT again to a final 1 mM concentration to further reduce residual disulfide bonds in peptides for 30 min.

  5. Add freshly prepared 1 M IAA to a final 10 mM concentration for 30 min in the dark to alkylate cysteine residues.

  6. Quench the unreacted IAA by adding 1 M DTT to 30 mM and incubate for 30 min.

  7. Desalt ~1 μg of each sample by C18 ZipTip according to the manufacturer’s protocol to test the digestion efficiency. Analyze each sample by LC-MS/MS.

  8. Perform a database search for MS raw data to check miscleavages by the percentage of identified miscleaved peptides. If the percentage is higher than 15% (see Note 11), add additional trypsin to further digest the samples.

  9. Acidify the samples by adding TFA to 0.5% (v/v). Check the pH by applying a small drop of the solution in a pH strip to ensure the pH is lower than ~3. Centrifuge at 21,000 × g for 10 min and take the supernatants.

  10. Use Ultra MicroSpin C18 desalting columns (cartridge) to desalt each sample as below.

  11. Wash the columns with 10 bed volumes of methanol and the elution buffer by centrifuging at 500 × g for 30 s (see Note 12).

  12. Equilibrate the columns with 10 bed volumes of the equilibration buffer.

  13. Load samples on the pre-equilibrated C18 columns by spinning at 100 × g for 3 min. Confirm that all the solution passes through the column. Reload the sample flow through once.

  14. Wash the columns with at least 10 bed volumes of the wash buffer.

  15. Elute the peptides with 5 bed volumes of the elution buffer twice at 100 × g for 5 min.

  16. Dry the eluted peptides by SpeedVac concentrator and store the peptides at −80 °C.

3.3. TMT16 Labeling of Peptides, Label Efficiency Test, Sample Pooling, and Labeled Peptide Desalting

For consistent quantification results, TMT labeling efficiency should be evaluated to confirm complete labeling for all samples. This step is vital because insufficient TMT reagents, inappropriate pH, and/or improper protein quantitation may lead to inefficient labeling. Additionally, a premix ratio test with a small aliquot of each sample is included to adjust the accurate mix ratio before pooling. The TMT16 reagents are used in this protocol.

  1. Resuspend each desalted peptide sample in 50 μL of 50 mM HEPES (pH 8.5) buffer, examine the pH using a pH strip paper to make sure the pH >8. Preserve ~1 μg of each unlabeled sample for TMT labeling efficiency test (see Note 13).

  2. Reconstitute TMT16 reagents in anhydrous acetonitrile according to the manufacturer’s instructions, add TMT16 reagent to each sample at a TMT/peptide ratio of 2:1 (w/w), and incubate at 21 °C for 30 min (see Note 14). Then save at −80 °C without quenching the reaction.

  3. Perform labeling efficiency test for each sample by LC-MS/MS with a short LC gradient: desalt ~1 μg of unlabeled and labeled samples using C18 ZipTips, and run LC-MS/MS for comparison. If complete TMT labeling is achieved, the unlabeled peptide peaks should not be found in the labeled sample data (see Note 15).

  4. Estimate the labeling efficiency by analyzing the MS1 intensity of the dominant, unlabeled peptides between unlabeled and labeled samples. For complete labeling, the unlabeled peptides should not be detected in the labeled samples (see Note 16).

  5. After ensuring complete labeling, quench the reaction for 15 min at 21 °C by adding 5% hydroxylamine to a final 1% concentration.

  6. Perform premix ratio test: mix 1 μL of each TMT-labeled sample. Confirm the accuracy of the pipettor before use. Desalt and analyze the mix by LC-MS/MS to determine the first mix ratio of each sample.

  7. Based on the first mix ratio (see Note 17), pool about half volume of each sample to make a mixture. Analyze 1 μg of aliquot of this mixture to obtain the second mix ratio of each sample.

  8. Based on the second mix ratio, add in the remaining samples to make an equal mixture (see Note 18).

  9. Acidify the pooled TMT-labeled samples by adding 10% TFA to pH < 3. Partially dry the sample to reduce ACN (reduce the volume to 70%). Centrifuge the pooled sample at 21,000 × g for 10 min and take the supernatant.

  10. Desalt the pooled samples by a C18 cartridge (see Subheading 3.2.2) to remove TMT-quenched byproducts, except that a more stringent wash condition is applied (the wash buffer of 0.1%TFA with 5% ACN for 10 bed volumes, see Note 19).

  11. Dry the eluted peptides in a SpeedVac concentrator and store the peptides at −80 °C.

3.4. Offline Basic pH RPLC Fractionation

The complexity of the CSF proteome due to extremely high dynamic range raises a challenge that is alleviated by extensive LC pre-fractionation of the CSF peptide samples before standard acidic pH RPLC-MS/MS analysis. Several pre-fractionation approaches have been developed, such as strong cation exchange (SCX), hydrophilic interaction chromatography (HILIC), and basic pH RPLC. The basic pH RPLC has the major advantage of high-resolving power, and this is used in this platform [35].

  1. Set up an XBridge C18 column containing bridged ethylene hybrid particles (3.5 μm particle size, 4.6 mm × 25 cm) driven by a HPLC system (e.g., Agilent 1220 infinity).

  2. Install a 100 μL loop and wash the loop with 300 μL of methanol, water, and buffer A using 3 loop volume. Wash the column with 100 μL washing solution (equally mixed isopropanol, methanol, acetonitrile, and water) and equilibrate the column in 95% buffer A for 1 h with a flow rate of 0.4 mL/min.

  3. Resuspend the pooled and desalted TMT16-labeled sample in 70 μL buffer A (see Note 20). Verify the sample pH value and adjust to pH 8.0 if necessary by adding 28% ammonium hydroxide (NH4OH). Centrifuge at 21,000 × g for 10 min to clarify the sample.

  4. Inject the sample and fractionate using the following gradient: 5% buffer B for 10 min, 5–15% buffer B for 2 min, 15–50% buffer B for 210 min, and 50–95% buffer B for 8 min, at the flow rate of 0.2 mL/min.

  5. For whole-proteome samples, a total of ~100 concatenated fractions are collected by fraction collector. Set the fractions collected every half minute.

  6. The concatenated fractions are dried in SpeedVac concentrator and stored at −80 °C for further LC-MS/MS analysis.

3.5. Acidic pH RPLC-MS/MS Analysis

The quantification is based on the TMT reporter ion fragmented during MS/MS or MS3 analysis. Ratio compression often occurs due to co-elution of TMT-labeled ions, which lead to co-isolation and co-fragmentation of these interfering ions [36]. Numerous strategies have been reported to address this ratio compression issue, including extensive fractionation [19, 37], narrow ion-isolation window [38], gas-phase separation [39], MultiNotch MS3 method [40], and computational correction [19]. To sufficiently resolve near-isobaric TMT16 reporter ions (6.32 mDa difference), we use 60,000 MS/MS resolution at 400 m/z. TMT reporter ion fragmentation is performed by higher energy collision dissociation (HCD), while electron transfer dissociation (ETD) is not recommended for this protocol, because ETD cleavage produces overlapping reporter ions. Collision-induced dissociation (CID) in ion trap is not possible for TMT-based quantification because of the 1/3 rule low mass limitation [41].

  1. Prepare a nanoscale column with C18 beads (1.9 μm beads, 75 μm ID with 15 μm tip orifice, and bed volume of 0.6 μL). Heat the columns at 65 °C by a butterfly portfolio heater to reduce backpressure.

  2. Wash the column with 95% buffer B and equilibrate column in 95% buffer A for at least 10 bed volumes.

  3. Check the LC-MS/MS system performance by running 100 ng of rat brain peptides or BSA peptides before analyzing the real samples. The following parameters need to be highly monitored for system evaluation: buffer levels, LC system pressure, survey MS signal intensity, quality of MS/MS spectra, mass shift, elution profile, and peak width (see Note 21).

  4. Dissolve the dried peptides from the basic pH RPLC in 5% formic acid. Centrifuge at 21,000 × g for 10 min and transfer the supernatant to an insert of a loading vial. For whole-proteome analysis, load 0.1–1 μg peptides for each fraction. Elute peptides by a gradient of 15–45% buffer B in 90 min (~0.25 μL/min flow rate). The loading amount and gradient may be adjusted to achieve the optimal results (see Note 22).

  5. The mass spectrometer (e.g., Q Exactive HF) is operated in data-dependent mode with a survey scan in Orbitrap (450–1600 m/z, 60,000 resolution, 1 × 106 automatic gain control (AGC), ~50 ms maximal ion time) and perform 20 data-dependent MS/MS high-resolution scans (60,000 resolution, 1 × 105 AGC target, ~150 ms maximal ion time, 32% HCD normalized collision energy (NCE), 1.0 m/z isolation window, 0.2 m/z isolation offset, and 10 s dynamic exclusion) (see Note 23).

3.6. MS Data Analysis

The computational pipeline of MS data processing includes searching protein databases for spectrum–peptide matches (PSMs), PSM filtering, and peptide/protein quantification. We have established a JUMP software suite to integrate all these steps [19, 29, 30] (see Note 24). The JUMP has been optimized for TMT-based analysis, including peptide identification and quantification, as well as the utilization of high-performance computing clusters to accommodate deep proteome analysis.

3.6.1. Database Search

The data analysis is performed using tag-based hybrid search engine JUMP software suite which comprises a hybrid database search engine (pattern- and tag-based). Briefly, the program converts MS raw files to mzXML format, preprocesses precursor ions to define charge state, generates peptide tags from MS/MS spectra, performs MS/MS pattern matching with theoretical patterns in the database, and finally outputs PSM ranking scores (Jscore). A non-redundant protein database is constructed (see Note 25). Searches are performed by allowing a 10 ppm mass tolerance for precursor ions, 15 ppm for product ions, and specifying additional parameters such as fully tryptic digestion with two maximal missed cleavages, three maximal modification sites per peptide and the assignment of a, b, and y ions. Static modifications include TMT16 tags on Lys residues and N termini (+304.20715 Da), and carbamidomethylation of Cys residues (+57.02146 Da), while a dynamic modification includes Met oxidation (+15.99491 Da).

3.6.2. Peptide–Spectrum Match Filtering

PSMs are filtered by JUMP-based matching scores (Jscore and ΔJn) and mass accuracy of precursor ion to reduce protein FDR to less than 1%. The resulting PSMs are grouped by peptide length (at least 7 amino acids), precursor ion charge, tryptic ends, modifications, missed cleavage sites, and charge state, and then filtered by matching scores to achieve defined FDR (see Note 26). The target-decoy strategy is used to evaluate FDR [42, 43]. For the peptides shared among multiple proteins (e.g., a protein family), the protein with the highest PSMs is selected to represent these shared peptides according to the rule of parsimony.

3.6.3. TMT-Based Protein Quantification

Quantification of peptides/proteins are achieved by averaging TMT reporter ion intensities of all matched PSMs. Briefly, from each PSM, TMT reporter ion intensities are extracted and corrected based on isotopic distribution of labeling reagents (e.g., 126 reporter ion produces 92.6%, 7.2%, and 0.2% of 126, 127C, and 128C m/z ions, respectively). If assuming equal protein amounts in all samples, the loading bias is corrected by mean or median intensities of all PSMs. Then each protein is quantified by averaging the quantitative data of all matched PSMs.

The quantification interference is corrected by using a previously described post-MS computational approach [19]. This y1 ion-based correction approach assumes that the y1 ion intensity is correlated to the reporter ion intensity. By deriving the linear relationship between y1 and reporter ion intensities from clean scans, the interference level in noisy scans is derived and corrected from the contaminated y1 ion intensity (see Note 27).

Finally, the peptide/protein quantification values are exported to a spreadsheet. Unsupervised data analysis methods, such as PCA or clustering analysis, are used to explore the relationship of samples. Differentially expressed proteins are identified by the statistical methods like t-test and analysis of variance (ANOVA).

Acknowledgments

We thank all other lab and center members for discussion and technical support. This work was partially supported by National Institutes of Health grants R01AG047928, R01AG053987, R01AG068581, RF1AG064909, U54NS110435, and American Lebanese Syrian Associated Charities (ALSAC).

Footnotes

1.

The JUMP software has been developed in house and optimized for TMT 16-plex analysis. Commercial software (e.g., Bioworks) may also be used for data processing.

2.

A highly active compound ammonium cyanate can be produced from dissolved urea which leads to carbamylation of amine groups in proteins; heating can accelerate this reaction. Hence, we use fresh urea solution for lysis and perform digestion at 21 °C to minimize the side reaction.

3.

Freshly made IAA solution is prepared and used at 21 °C to avoid the side reaction with Lys residues [44]. Modification of Lys residues occurs at a higher temperature to form a single (57.02146 Da) and double tag (114.04293 Da, identical mass as the GG tag generated by tryptic digestion of ubiquitin) [45].

4.

In this protocol, to prevent autolysis of trypsin, the enzyme is acetylated at the ε-amino group of lysine residues. The main reasons to use the modified trypsin are: (a) unmodified trypsin undergoes auto-proteolysis to generate peptides that affect MS analysis; and (b) trypsin auto-proteolysis produces pseudo trypsin that has chymotrypsin-like activity.

5.

The selection of desalting C18 columns is based on the input sample amount. Excessive or limited bed volumes result in low recovery of peptides. Cartridge capacities are ~3–5% of the sorbent weight [46], or 10 μg of peptides per μL of C18 resin [47].

6.

The ideal sample loading amount differs on various LC systems. We usually load less than 3 μg of peptides per μL of C18 resin for peptide separation and perform a pilot experiment to test the LC system before running real samples [33]. HPLC operation pressure is another factor that needs to be considered, since it is determined by column internal diameter, length, resin particle size, resin porous feature, and buffer viscosity.

7.

The addition of dimethyl sulfoxide in the LC buffers improves peptide ionization efficiency to enhance sensitivity, although it also increases background ions to affect long-term performance of MS instruments. Frequent MS cleaning and maintenance may be required [48].

8.

Protein concentration can be measured by the BCA assay or short SDS gel staining method. Although the BCA assay is commonly used, we suggest to validate some of the sample concentrations by a short SDS gel because numerous non-protein constituents in the samples often lead to protein overestimation by BCA.

9.

The quality control step is critical before the TMT experiment. The long SDS-PAGE gel is highly recommended to examine the quality and exclude the degraded samples. For the samples with known protein expression profile, Western blotting can be performed for sample validation. This step may identify possible mislabeling or other issues during sample preparation.

10.

It is important to consider the CSF volume when calculating the final concentration of lysis buffer (e.g., in an Excel table).

11.

We usually observe that the percentage of peptides with miscleaved sites from mammalian protein mixtures is below 15%. The remaining miscleaved sites are usually due to surrounding acidic residues and/or ragged ends (e.g., KK, RR, KR, and RK).

12.

This step is used to wet the columns and elute possible contaminants on the columns.

13.

Incomplete drying of highly acidic peptide samples after desalting may lead to acidic pH, which inhibit TMT labeling. It is important to verify the pH ~8 by pH strips.

14.

Because the mass of TMT16 reagents is 1.2-fold larger than that of TMT11 reagents, the TMT16/protein ratio is also higher than TMT11/protein ratio.

15.

For the LC-MS/MS of TMT labeling efficiency test, blank runs can be performed to remove any carryover of unlabeled peptides from previous users in the HPLC system. We then run TMT-labeled samples before unlabeled samples. This order is important to avoid the unlabeled peptide carryover to contaminate the TMT-labeled samples.

16.

We often select 5–10 dominant peptides to confirm the labeling efficiency. If not fully labeled, additional TMT reagents are added to continue the labeling.

17.

The premix ratio test identifies the sample of the lowest amount, which is used as the baseline to adjust the amounts of other samples.

18.

Pipetting errors may affect the accuracy of protein/peptide quantitation. The variation among 16 samples is controlled to be less than 5–10% after multiple rounds of adjustment.

19.

The stringent wash condition is used for desalting TMT16-labeled sample as the derivatives in the TMT16 quenching reaction (TMT16-OH and TMT16-NHOH) are more hydrophobic than TMT11 counterparts.

20.

To reduce sample loss during loading, the sample volume is usually less than 70% of the loop volume.

21.

To improve the LC-MS/MS system performance, regular maintenance, cleaning, mass calibration, tuning of MS instrument, and preparing fresh LC buffers are highly recommended. Mass accuracy can be assessed by polysiloxane ion 445.12003.

22.

To obtain high identification number, run one of the fractions (often in the middle of basic pH RPLC) to test the loading amount and elution gradient. The gradient adjustment is based on the distribution of the identified peptides along the retention time. A proper gradient should result in evenly distributed peptides during elution.

23.

The parameters used are optimized based on our settings in Q Exactive HF. The optimal NCE settings for TMT11 and TMT16 are 35% and 32%, respectively.

24.

The data analysis is performed using JUMP software which involves a pattern-based database search and a tag-based de novo sequencing to improve sensitivity and specificity. Data analysis can also be performed with other commercially or freely available software.

25.

We generate the non-redundant database by combining protein sequences from Swiss-Prot, TrEMBL, and UCSC databases. One can also add customized protein sequences not present in those reference databases, including protease cleaved proteins, proteins with single-nucleotide polymorphisms, and other common contaminants.

26.

If positive-control peptides/proteins are missing at the filtering step, FDR and other parameters (e.g., peptide length) may be adjusted to recover the expected peptides and validate by manual examination.

27.

For TMT-labeled tryptic peptides, K-TMT and R residues are two symbolic y1 ions (376.27574 Da and 175.11895 Da, respectively) in MS/MS spectra. If only one y1 ion is detected in a spectrum and is consistent with the identified peptide, the spectrum is considered to be a clean scan. If both y1 ions are detected, the spectrum is regarded as a noisy scan.

References

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