A Comprehensive Workflow for Plasma Proteomics by Mass Spectrometry: A Strategy for Biomarker Discovery in Systemic Human Diseases

Summary

Plasma proteomics enables biomarker discovery for systemic diseases through minimally invasive sampling. Despite analytical challenges posed by plasma complexity, this study presents an optimized liquid chromatography-mass spectrometry (LC-MS/MS) workflow that enhances proteome coverage, reduces variability, and improves reproducibility, enabling accurate comparisons between disease and control samples.

Abstract

Plasma, the cell-free component of blood, offers a minimally invasive and rich source of biomolecular information, making it ideal for investigating systemic diseases. The advent of high-throughput proteomic technologies has revolutionized plasma analysis by enabling large-scale identification and quantification of circulating proteins. These advances have facilitated biomarker discovery for early diagnosis, prognosis, and therapeutic targeting of various disorders, including cancers and inflammatory diseases. However, the inherent complexity of plasma, along with inconsistent sample preparation and analysis methods, has historically limited proteome coverage and reproducibility. Early approaches relying on crude plasma analysis often suffered from poor consistency and significant batch effects, impeding reliable comparative studies across clinical cohorts.

To overcome these limitations, the current study introduces a robust and scalable plasma proteomics workflow. The protocol integrates key preparatory steps such as immunodepletion, molecular weight cutoff filtration, lyophilization, protein quantification and normalization, enzymatic digestion, and LC-MS/MS profiling. This optimized strategy significantly enhances proteome depth and reduces variability, thereby enabling more accurate and reproducible differential protein expression analysis between disease cases and healthy controls. The workflow provides a valuable platform for researchers aiming to generate high-quality proteomic data from plasma, ultimately contributing to the advancement of biomarker-driven diagnostics and personalized medicine.

Introduction

Plasma, the cell-free component of blood, is gaining increasing recognition as a reservoir of biological information relevant for disease studies. It carries a plethora of diverse circulating biomolecules, ranging from proteins and metabolites to lipids and nucleic acids, which is a mirror of the organism's physiological and pathological status. This systemic nature of plasma makes it an ideal biospecimen for investigation into disease pathophysiology, ranging from cancer to metabolic and inflammatory disorders. A major benefit of using plasma as a sample lies in its minimally invasive collection method, allowing for repeated sampling in clinical settings, which is important for observation of disease progression or therapeutic response. However, plasma exhibits an extremely complex proteomic matrix, whereby proteins such as albumins and immunoglobulin levels span more than ten orders of magnitude, masking identification¹.

Over the past two decades, plasma proteomics has enabled the identification of clinically relevant biomarkers such as beta-human chorionic gonadotropin (β-hCG) and troponin I, driven by advances in large-scale protein analysis technologies². These tools accept, as input, highly sensitive information, and with their high-throughput nature, provide large-scale profiling of plasma proteins. With the advent of liquid chromatography-mass spectrometry (LC-MS/MS) platforms, it is now possible to analyze thousands of protein species using as little as 10-50 µL of plasma input, with a dynamic detection range spanning several orders of magnitude^3,4. This has increased the use of plasma proteomics in translational research: early disease detection, improved prognostic models, and the discovery of new therapeutic targets⁵.

An important milestone in this area was the establishment of the Human Plasma Proteome Project (HUPO-HPPP), which systemically catalogued plasma proteins and demonstrated the feasibility of plasma proteomics as an established field⁶. Afterwards, the Clinical Proteomic Tumor Analysis Consortium (CPTAC) adopted rigorously validated protocols and cross-laboratory studies to address issues of reproducibility that earlier confounded proteomic research⁷. These initiatives underscored the need for standardization and highlighted key procedures, including uniform plasma handling, depletion of abundant proteins, enzymatic digestion, and consistent LC-MS/MS parameters. Recent studies by Geyer and colleagues showed that plasma proteomic signatures can be reliably generated across many individuals when standardized methodologies are used, providing a basis for population-level studies and biomarker validation processes⁸.

The progression of plasma proteomics has been shaped considerably by the recognition of quality control (QC) and performance metrics as critical components of experimental design. Conventionally, differences in digestion efficiency, peptide identification rates, and instrument performance have led to inconsistencies across research studies, thereby affecting reproducibility. Many studies have been undertaken to address the issues. Gawor et al. outlined measurable quality indicators for sample preparation and mass spectrometry performance, thereby providing a systematic method for assessing the robustness of the protein profiling pipeline⁹. Tsantilas et al. presented computerized benchmarking tools that allow researchers to compare datasets across platforms and laboratories, thereby increasing transparency in the presentation of proteomics data¹⁰. Moreover, advances in methodology, such as immunodepletion, size-exclusion fractionation, and Data-independent acquisition (DIA), have enhanced proteome coverage and reduced technical bias^11,12. Such quality assurance measures ensure that plasma proteomic data are reproducible and serve as a scaffold for future clinical translation studies.

Despite these significant advances, plasma's inherent complexity continues to harbor challenges. It is common for direct analysis of crude plasma to yield suboptimal proteome coverage, as very abundant proteins overwhelm low-abundance analytes, many of which are the most promising biomarker candidates. Moreover, technical variation stemming from variable protein quantitation, incomplete digestion, and inadequate cleanup can contribute to batch artifacts and decreased cohort comparability¹³. These limitations underscore the need for highly optimized workflows that balance proteome depth, reproducibility, and scalability. This study protocol addresses this need by demonstrating an optimized plasma proteomics workflow that combines immunodepletion, molecular-weight-cutoff filtration, lyophilization, protein normalization and quantitation, enzymatic digestion, and LC-MS/MS profiling. By increasing proteome coverage and reducing technical variation, the methodological framework enables consistent relative quantitation between disease-affected and control groups. By introducing scalability and accommodating downstream analyses, the workflow provides a usable, reproducible method for extracting clinically significant information from plasma proteomics.

Protocol

This protocol outlines a streamlined workflow designed to enhance plasma proteomics for biomarker discovery in systemic inflammatory diseases^14,15. The study was done in accordance with the Institutional Ethics Committee and the Helsinki Declaration of 1975, as revised in 2000. Informed consent was obtained from the participants. Initially, plasma samples undergo immunodepletion to remove high-abundance proteins, followed by molecular weight-based filtration to reduce sample complexity. The samples are then lyophilized and quantified to ensure protein concentration is adequate for downstream processing. After normalization, proteins are subjected to tryptic digestion to generate peptides suitable for analysis. Finally, mass spectrometry is employed to detect and quantify low-abundance proteins, facilitating reliable comparisons between healthy and disease conditions (Figure 1) shows the blood proteomics pipeline.

1. Sample collection:

1. Obtain 3 mL of blood from participants in Ethylenediaminetetraacetic acid (EDTA) Vacutainer tubes by venipuncture from the antecubital vein under aseptic conditions. This is done by a trained phlebotomist.
2. To separate the plasma, centrifuge the samples (fixed-angle rotor with 12 × 15 mL ) for 10 min at 704 g. Keep the plasma samples in storage at -80 °C until subsequent examination.

2. Immunodepletion

NOTE: Immunodepletion is an affinity chromatography-based method used to selectively remove high-abundance proteins from biological fluids like serum, plasma, and cerebrospinal fluid (CSF). A spin cartridge containing immobilized antibodies targets and captures 14 common high-abundance proteins, including albumin, immunoglobulins (IgG, IgA, IgM), transferrin, and complement C3. These proteins are retained in the cartridge while low-abundance proteins pass through for further analysis. The technique enhances proteomic profiling by reducing sample complexity and improving the detection of potential biomarkers. This approach is widely adopted in plasma proteomics to improve sensitivity and reproducibility^16,17.

1. Cartridge preparation
   1. Remove the 0.45-mL spin cartridge from the refrigerator. Leave it to equilibrate to ambient temperature.
   2. Prepare Buffer A Equil/Load/Wash and Buffer B Elution according to the number of samples processed. Apply approximately 5 mL of Buffer A and 2 mL of Buffer B per 10 µL plasma sample. Decant into 50 mL sterile centrifuge tubes.
   3. Label two sterile Luer-Lok 5 mL syringes as A for Buffer A and B for Buffer B.
2. Plasma sample preparation
   1. Dilute each 10 µL of plasma sample with 190 µL of Buffer A to a final volume of 200 µL.
      NOTE: Add protease inhibitors to Buffer A to avoid degradation of plasma proteins while processing.
   2. A plasma sample may contain particulate matter. It may clog the cartridge. In order to prevent clogging and improve the sample clarity, centrifuge the diluted sample through a 0.22 µm spin filter for 1 min at 16,000 ×g at 4 °C.
3. Cartridge setup
   1. Remove the cartridge cap and the bottom cap and store them for use later.
   2. Place a Luer-Lok adapter on the spin cartridge.
   3. Pull 4 mL of Buffer A in the syringe labeled "A" and connect the syringe to the Luer-Lok adapter.
   4. Gradually flow Buffer A through the cartridge to allow the resin to equilibrate and displace any entrapped air.
   5. Aspirate off the excess buffer from the top of the cartridge with a sterile micropipette.
   6. Remove the Luer-Lok adapter and deposit the spin cartridge into a screw-cap collection tube marked F1.
4. Sample loading and fraction collection
   1. Add 200 µL of filtered, diluted plasma directly onto the resin bed.
   2. Centrifuge the cartridge at 100 × g for 1 min at 4 °C. Leave the cartridge uncapped or loosely capped to allow smooth flow.
   3. Collect the flow-through fraction F1 in the labeled collection tube. Ensure the resin bed and frit remain moist after centrifugation.
   4. Let the cartridge stand at room temperature for 5 min.
5. Washing and recovery of flow-through fractions
   1. Add 400 µL of Buffer A onto the resin bed and centrifuge at 100 × g for 2.5 min at 4 °C. Add this eluate to the F1 collection tube.
   2. Transfer the cartridge to a new collection tube labeled F2.
   3. Add another 400 µL of Buffer A and centrifuge at 100 × g for 2.5 min at 4 °C. Collect the eluate in the F2 tube.
6. Elution of bound proteins
   1. Remove the cartridge from the F2 tube and attach the Luer-Lok adapter.
   2. Fill the syringe labeled "B" with 2.5 mL of Buffer B and attach it to the cartridge.
   3. Slowly push Buffer B through the cartridge to elute the bound high-abundance proteins into a fresh collection tube.
      NOTE: Save the bound fraction if further analysis is required; otherwise, discard appropriately.
7. Cartridge re-equilibration
   1. Remove the syringe containing Buffer B from the cartridge and attach the syringe labeled "A" containing 5 mL of Buffer A.
   2. Slowly push Buffer A through the resin bed to re-equilibrate the cartridge at a flow rate of 0.5 mL per minute.
      NOTE: Do not allow the resin bed or frit to dry. Leave a small amount of buffer on top of the frit during storage.
   3. Once equilibrated, the cartridge is ready for the next sample.
8. Downstream use
   1. For maximum recovery of low-abundance proteins, combine F1 and F2 flow-through fractions.
      ​NOTE: F1 and F2 may also be analyzed separately if differential analysis is required. The buffer added is 1 mL, so the combined F1 fraction may be ~850 µL to ensure good recovery. Analyse the crude plasma, flow-through, and eluate by Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) to confirm removal of the top 14 high-abundance proteins from the flow-through. 12.5 µL of flow through F1, crude plasma, and eluate will be used for loading in gel to make a well volume of 25 µL (Laemmli sample buffer: sample ratio 2:1)

3. Protein concentration using a 5 kDa spin concentrator

NOTE: Concentrating a sample using a 5 kDa spin concentrator is based on the principle of size exclusion and molecular filtration to achieve rapid concentration and high recovery of target biomolecules in a single tube. This enables concentration of proteins by removal of solvent and buffer exchange or desalting of the protein solution.

1. Fill the spin concentrator with up to 4 mL of the protein sample.
2. Place the spin concentrator into a refrigerated centrifuge.
3. Centrifuge at 5,000 × g at 10 °C for 30 min, or until the desired volume reduction (e.g., 30× concentration) is achieved.
4. Remove the spin concentrator from the centrifuge and recover the concentrated sample from the bottom of the retentate chamber using a micropipette. To ensure good recovery, expect ~150 µL concentrate recovered from a total of 850 µL protein sample.
   NOTE: Ensure that the membrane does not dry out during centrifugation to maintain protein integrity and recovery efficiency.

4. Desalting and buffer exchange

1. Discard the filtrate from the bottom chamber of the concentrator.
2. Refill the centrifuge tube with an equal volume of 50 mM ammonium bicarbonate buffer (pH 7.2-7.4). (For example, if the retentate is 150 µL, add 150 µL of buffer)
3. Centrifuge again at 5,000 × g at 10 °C until the original concentration is achieved.
4. Repeat this buffer exchange process three times to remove approximately 99% of small molecule contaminants such as salts or urea.
5. Concentrate the protein samples by freeze-drying them using a speed vacuum lyophilizer.
   NOTE: Adjust the number of cycles based on the degree of desalting required for downstream applications.

5. Protein quantification using Bicinchoninic acid (BCA) assay

NOTE: The protein concentration of the samples was determined using the BCA protein assay, with bovine serum albumin (BSA) as the standard. The BCA assay is a detergent-compatible, bicinchoninic acid-based colorimetric method for detecting and quantifying total protein. It relies on the reduction of Cu²⁺ to Cu⁺ by proteins in an alkaline environment (the Biuret reaction), followed by highly sensitive and selective colorimetric detection of the cuprous ion (Cu⁺) using a reagent containing bicinchoninic acid. The assay produces a purple-colored complex formed by the chelation of one cuprous ion with two molecules of BCA. The BCA assay produces a consistent, linear response with BSA, ensuring a reliable standard curve reference. Although it is known that different proteins yield different responses in the BCA assay, using BSA provides reproducible relative quantitation across diverse samples. Since our aim was normalization for equal protein loading rather than absolute quantification of every species, BSA-based calibration offered the most robust and reproducible approach.

1. Procedure
   1. Add 25 µL of each BSA standard (working range 20-2000 µg/mL) and the unknown protein sample into designated wells of a 96-well plate.
   2. Add 200 µL of BCA working reagent to each well.
   3. Mix thoroughly using a microplate shaker for 30 s.
   4. Cover the plate and incubate at 37 °C for 30 min.
   5. Bring the plate to room temperature.
   6. Measure absorbance at 562 nm using a microplate reader.
2. Data analysis
   1. Subtract the average absorbance of the blank (zero protein) from all sample and standard absorbance values.
   2. Plot a standard curve of absorbance vs. BSA concentration (microgram/microliter).
   3. Use the regression equation from the curve to calculate protein concentrations in unknown samples.

6. Normalization of protein concentration

1. Procedure
   1. Measure the protein concentration of each plasma sample using an appropriate quantification method.
   2. Record the concentration and total volume of each sample.
   3. Prepare 50 mM ammonium bicarbonate buffer (pH 7.8) as the diluent.
   4. Calculate the required dilution using the formula C₁V₁ = C₂V₂.
   5. For a sample with a concentration of 1.3 mg/mL and a volume of 100 µL, calculate the final volume required to reach 1 mg/mL:[V₂ = \frac{1.3 \times 100}{1} = 130\ \mu L]
   6. Calculate the volume of buffer to add by subtracting the initial volume from the final volume:Buffer volume = 130 µL − 100 µL = 30 µL.
   7. Add 30 µL of 50 mM ammonium bicarbonate (pH 7.8) to the 100 µL plasma sample.
   8. Mix the sample gently by pipetting or brief vortexing.
   9. Confirm that the final volume is 130 µL at a normalized concentration of 1 mg/mL.
   10. Proceed with downstream proteomic sample preparation.
       NOTE: Ensure accurate concentration adjustment by calculating the required dilution factor based on the results of the BCA assay. Each step is validated, and the data is given as supplementary data (Supplementary Table 1).

7. Sample preparation and tryptic digestion

1. Protein reduction
   1. Add an appropriate reducing agent (e.g., 1,4-dithiothreitol [DTT]) to the protein solution in ammonium bicarbonate buffer to a final concentration of 100 mM.
   2. Incubate the mixture at 60 °C for 30 min in a thermomixer or heating block.
      CAUTION: DTT is an irritant and should be handled with gloves inside a fume hood.
      ​NOTE: Additional information on the trypsin used for digestion has been provided in the Table of Materials. Briefly, sequencing-grade Tosyl Phenylalanyl Chloromethyl Ketone (TPCK)-treated trypsin (to eliminate chymotryptic activity) was used to ensure high specificity for cleavage at lysine and arginine residues. The enzyme activity was verified based on manufacturer specifications, corresponding to a N-Benzoyl-L-Arginine Ethyl Ester (BAEE) activity of approximately 1 unit per mg of protein. Regarding the DTT and Iodo acetamide (IAA) concentrations, we confirm that the 100 mM DTT and 200 mM IAA refer to stock solutions, not the final concentrations used in the reaction. The final working concentrations during reduction and alkylation were within the standard range (typically 5-10 mM DTT and 10-20 mM IAA, respectively).
2. Alkylation of cysteine residues
   1. Allow the sample to cool at room temperature for 5 min after reduction.
   2. Briefly centrifuge to collect condensate at the bottom of the tube.
   3. Add 200 mM IAA prepared in 50 mM Ammonium Bicarbonate (ABC) buffer to alkylate the reduced cysteine residues.
   4. Incubate the sample in the dark at room temperature for 30 min to prevent IAA degradation.
      ​NOTE: Iodoacetamide is light-sensitive and potentially toxic; avoid exposure and handle in subdued light.
3. Enzymatic digestion
   1. Prepare sequencing-grade modified trypsin in 50 mM ABC buffer immediately before use.
   2. Add trypsin to each sample at a trypsin-to-protein ratio of 1:25 (w/w).
   3. Incubate the reaction at 37 °C for 17 h to allow complete enzymatic digestion.
      ​NOTE: Extended digestion times may lead to overdigestion; ensure temperature stability throughout incubation.
4. Quenching the reaction
   1. Stop the enzymatic reaction by adding formic acid to each sample to achieve a final concentration of 1.0% (v/v).
   2. Incubate at 37 °C for 20 min to denature residual trypsin and stabilize the peptide mixture.
      ​CAUTION: Formic acid is corrosive; use appropriate personal protective equipment (PPE) and work in a chemical fume hood.
5. Peptide recovery
   1. Centrifuge the digested peptide mixture at 3500 g for 12 min at room temperature to remove insoluble material.
   2. Carefully transfer the clear supernatant containing digested peptides into autosampler vials without disturbing the pellet.
   3. Store the peptide solutions at -20 °C until further LC-MS/MS analysis.
6. Preparation for LC-MS analysis
   1. Load the peptide samples directly onto the LC-MS system equipped with ion mobility for peptide separation and analysis.
   2. Ensure that all vials are properly sealed to prevent evaporation and contamination.

8. LC MS-MS analysis

1. Peptide loading and desalting
   1. Load 1 µg of the peptide sample into a sample loop in partial-loop injection mode.
   2. Desalt peptides using a C18 trap column by washing with aqueous solvent containing 0.1% formic acid at a flow rate of 15 µL/min for 1 min.
      NOTE: Use water with 0.1% formic acid as Solvent A and acetonitrile with 0.1% formic acid as Solvent B throughout the run.
2. Peptide separation
   1. Connect the trap column in-line with an analytical reversed-phase C18 column.
   2. Elute peptides using a linear gradient from 3% to 40% Solvent B over 55 min at a flow rate of 300 nL/min.
   3. Rinse the column with 80% Solvent B for 7.5 min to remove retained components.
   4. Re-equilibrate the column with 3% Solvent B before the next injection.
   5. Maintain the column oven at 35 °C and the autosampler at 4 °C during the run.
      NOTE: Include blank injections between samples to minimize carryover.
3. Mass spectrometry acquisition settings
   1. Mass spectral analysis of eluting peptides
      1. Set up the UPLC system and couple it directly to the High-Definition MS.
      2. Control the system using MS software.
      3. Ensure that the mass spectrometer is configured as a hybrid quadrupole-ion mobility-orthogonal acceleration-time-of-flight (oa-TOF) instrument.
      4. Verify that all nitrogen and lock-spray gas connections are secure and leak-free before initiating operation.
         CAUTION: Handle high-voltage connections and pressurized gases in compliance with laboratory safety standards.
   2. Ionization setup
      1. Operate the mass spectrometer in positive electrospray ionization (ESI) mode using a dual electrospray ion source.
      2. Set the nano-ESI capillary voltage to 3.4 kV.
      3. Adjust the sample cone voltage to 40 V and the extraction cone voltage to 4 V.
      4. Set the ion mobility spectrometry (IMS) nitrogen gas flow to 90 mL/min.
      5. Inspect the spray stability and ensure consistent ion current before proceeding.
   3. Ion mobility separation
      1. Set the IMS ion guide pulse height to 40 V during ion transmission.
      2. Adjust the IMS ion guide velocity to 800 m/s.
      3. Ramp the traveling wave height linearly from 8 V to 20 V over the complete IMS cycle to achieve optimal ion separation by mobility.
         CAUTION: Ensure stable IMS gas flow; fluctuations may affect ion separation and drift time reproducibility.
   4. Lock mass calibration
      1. Calibrate the time-of-flight (TOF) analyzer using a 2 µg/µL sodium iodide (NaI) solution.
      2. Set the calibration range to detect ions between 50 m/z and 2000 m/z.
      3. Acquire lock mass reference spectra every 45 s using the dual electrospray ion source channel for real-time mass correction.
      4. Verify that the calibration curve achieves mass accuracy within ±5 ppm before proceeding to data acquisition.
4. Data acquisition
   1. Acquire data in continuum mode, alternating between two functions:
      1. Function 1 (Low Energy MS): Record low-energy precursor ion spectra.
      2. Function 2 (High Energy HDMSE): Record fragment ion spectra at elevated collision energy with ion mobility separation.
   2. In Function 2, set the collision energy in the Trap region to 4 eV.
   3. Ramp the collision energy in the Transfer region from 20 eV to 45 eV to achieve controlled peptide fragmentation.
   4. Set the spectral acquisition time per function to 0.9 s and the interscan delay to 0.024 s.
   5. Monitor the total ion chromatogram (TIC) to confirm consistent instrument response throughout the run.
   6. Import raw data into proteomics analysis software (e.g., Progenesis QI or equivalent).
   7. Align chromatograms across runs using the default alignment reference or a manually selected reference run.
   8. Identify peptides and proteins using a curated database (e.g., UniProt Human - reviewed entries only).
   9. Set the following search parameters:
      Enzyme: Trypsin
      Fixed modification: Carbamidomethylation (C)
      Variable modification: Oxidation (M)
      Peptide tolerance: According to the instrument resolution
      Missed cleavages allowed: 1
      False Discovery Rate (FDR): ≤ 1%
   10. Perform normalization using the normalize to all proteins method.
   11. Apply filtering thresholds: ≥2 peptides per protein, ≥1 unique peptide, fold change ≥1.5, and adjusted p-value <0.05.
       NOTE: This is general laboratory safety guidance-always follow the institution's EHS (Environmental Health & Safety) regulations and local legal requirements.
5. Quality control
   1. Perform pre- and post-run calibration checks using the NaI standard to confirm mass accuracy.
   2. Ensure that the lock mass reference remains stable throughout the analysis.
   3. Evaluate replicate injections to confirm reproducibility in retention time, drift time, and spectral intensity.
      CAUTION: Sodium iodide is an irritant. While handling, use gloves and avoid inhalation or contact with skin.
      NOTE: Refer to Supplementary File 1 for waste disposal guidance. Always work in a cold environment or on ice when handling plasma samples to prevent protein degradation. Wear Nitryl gloves, a lab coat, and eye protection while handling biological samples and buffers. Perform centrifugation steps using a refrigerated centrifuge pre-cooled to 4 °C. Ensure all centrifuge tubes and syringes used are sterile and properly labeled to avoid cross-contamination. Always use proteomic-grade water for preparing buffers.

Results

Plasma samples were processed using a spin cartridge to selectively remove 14 high-abundance proteins, namely albumin, IgG, IgA, transferrin, haptoglobin, antitrypsin, fibrinogen, alpha2-macroglobulin, alpha1-acid glycoprotein, IgM, apolipoprotein AI, apolipoprotein AII, complement C3, and transthyretin. This depletion step was critical to enhance the detection of low-abundance biomarkers in downstream proteomic analysis. Each plasma sample (8-10 µL) was diluted, clarified via 0.22 µm spin filtration, and passed through ...

Discussion

The intrinsic complexity of plasma as a biological matrix presents a major challenge to proteomic analysis. The enormous dynamic range of protein concentrations, along with the predominance of a few high-abundance species, can mask the detection of clinically relevant lower-abundance proteins. For this current study, we mitigated these disadvantages utilizing a multi-step, standardized pipeline for sample preparation that was designed to selectively enrich moderate- and low-abundance proteins and to maintain analytical c...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
5 kDa Molecular Weight Cutoff Spin Concentrator	Agilent Technologies	5185−5991	Used for buffer exchange and salt removal; retains proteins above 5 kDa
Ammonium Bicarbonate (ABC)	Sigma Aldrich	Cat no: T6567	Used for Tryptic digestion
Buffer A Equil/Load/Wash	Agilent Technologies	5185-5987
Buffer B Elution	Agilent Technologies	5185-5988
Cool centrifuge	CPR-30 Plus Refrigerated Centrifuge	N/A	Used for centrifuging protein samples at low temperature
DTT	Sigma Aldrich	CASNo: 3483-12-3	Used for Tryptic digestion
Electrophoretic apparatus	BIO RAD	1658004	Used for SDS PAGE
IAA	Sigma Aldrich	CAS No: 144-48-9	Used for Tryptic digestion
IMS T-Wave	Waters	https://www.waters.com/content/dam/waters/en/library/white-papers/2012/water-whitepapers-Triwav eMoreCompleteCharacterizationofMi xturesandMolecules-720004176.pdf	Ion mobility separation (IMS)
Infinite M Plex Multi mode reader	TECAN		Multiplate reader
LC-MS system	Waters	https://www.waters.com/nextgen/us/en/products/mass-spectrometry/mass-spectrometry-systems/liquid-chromatography-ms-systems.html
MassLynx 4.1 SCN781 software	Waters	https://www.waters.com/nextgen/us/en/products/informatics-and-software/mass-spectrometry-software/masslynx-mass-spectrometry-software.html	Mass spectrometry software
Multiple Affinity Removal Spin Cartridge Human 14 (MARS Hu-14)	Agilent Technologies	5188−6560	Used to deplete 14 high-abundance plasma proteins including albumin, IgG, IgA, transferrin, etc.
nanoACQUITY UPLC system	Waters	https://www.waters.com/nextgen/en/products/chromatography/chromatography-systems/acquity-uplc-m-class-system.html
NanoLockSpray	Waters	https://www.waters.com/nextgen/us/en/services/instrument-services/instrument-upgrades/mass-spectrometry-ms-upgrades/nanolockspray-exact-mass-ionization-source-upgrade.html
Operon FDU-7003	Operon	http://www.operon.co.kr/en/pro2-FDU.html	speed vacuum lyophilizer
Pierce BCA protein assay kit	Thermo Scientific	23225	Used to quantify protein concentration using colorimetric assay
Progenesis QI for Proteomics	Waters	V4.2
R-8C BL Laboratory centrifuge, fixed-angle rotor with 12 × 15 mL	REMI		Centrifuge
SYNAPT G2 high-definition MS	HDMSE System, Waters Corporation	https://www.waters.com/content/dam/waters/en/app-notes/2009/720003057/720003057-en.pdf	Hybrid quadrupole IMS orthogonal acceleration time-of-flight (oa-Tof) mass spectrometer
Total recovery vial	Waters	https://www.waters.com/nextgen/us/en/shop/vials-containers--collection-plates/186004631-clear-glass-12-x-32-mm-screw-neck-total-recovery-vial-preassembl.html
Trypsin	Sigma Aldrich	Cat no: T6567	Used for Tryptic digestion