"Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome

Summary

Here, we describe a proteomics workflow for characterization of the cell surface proteome of various cell types. This workflow includes cell surface protein enrichment, subsequent sample preparation, analysis using an LC-MS/MS platform, and data processing with specialized software.

Abstract

Over the past decade, mass spectrometry-based proteomics has enabled an in-depth characterization of biological systems across a broad array of applications. The cell surface proteome ("surfaceome") in human disease is of significant interest, as plasma membrane proteins are the primary target of most clinically approved therapeutics, as well as a key feature by which to diagnostically distinguish diseased cells from healthy tissues. However, focused characterization of membrane and surface proteins of the cell has remained challenging, primarily due to the complexity of cellular lysates, which mask proteins of interest by other high-abundance proteins. To overcome this technical barrier and accurately define the cell surface proteome of various cell types using mass spectrometry proteomics, it is necessary to enrich the cell lysate for cell surface proteins prior to analysis on the mass spectrometer. This paper presents a detailed workflow for labeling cell surface proteins from cancer cells, enriching these proteins out of the cell lysate, and subsequent sample preparation for mass spectrometry analysis.

Introduction

Proteins serve as the fundamental units by which the majority of cellular functions are carried out. Characterizing the structure and function of relevant proteins is an essential step to understand biological processes. Over the past decade, advances in mass spectrometry technology, analysis software, and databases have enabled the accurate detection and measurement of proteins at a proteome-wide scale¹. Mass spectrometry-based proteomics can be utilized in a diverse array of applications, from basic science analysis of biochemical pathways, to identification of novel drug targets in a translational setting, to diagnosis and monitoring of diseases in the clinic². When screening for novel drug targets, characterization of the cell surface proteome is particularly important, with over 65% of currently approved human drugs targeting cell surface proteins³. The field of cancer immunotherapy also wholly relies on cancer-specific cell surface antigens to target and specifically eliminate tumor cells⁴. Mass spectrometry-based proteomics can thus serve as a promising tool to identify new cell surface proteins toward therapeutic interventions.

However, there are several limitations when utilizing conventional proteomics methods to survey tumor cells for novel cell surface protein targets. A primary concern is that surface proteins make up a very small fraction of the total protein molecules in a cell. Therefore, fragments of these proteins are masked by a high abundance of intracellular proteins when performing mass spectrometry analysis of the whole-cell lysate⁵. This limitation makes it challenging to accurately characterize the cell surface proteome with a traditional proteomics workflow. To address this challenge, it is necessary to develop ways to enrich cell surface proteins out of the whole-cell lysate, prior to analysis on the mass spectrometer. One such method involves the oxidation and biotin labeling of glycosylated cell surface proteins in the intact cells, and subsequent enrichment of these biotinylated proteins from the lysate with a neutravidin pulldown, a process that has been termed "cell surface capture"⁶. Since ~85% of mammalian cell surface proteins are thought to be glycosylated⁷, this serves as an effective method of enriching the cell surface proteome out of the whole cell lysate. This paper describes a complete workflow, beginning with cultured cells, of cell surface biotin labeling, and subsequent sample preparation for mass spectrometry analysis (Figure 1). Over several replicates, this method provides robust coverage of the cell surface proteome of a particular sample. Utilizing this method to characterize the cell surface proteome of both tumor and healthy cells can facilitate the discovery of novel cell surface antigens to identify potential immunotherapeutic targets⁸.

Protocol

NOTE: AMO1 plasmacytoma cells were used for this cell surface proteome experiment. The same protocol could be used for other cell types as well, including a wide array of suspension and adherent cell lines⁹, as well as various types of primary samples¹⁰. However, cell numbers (starting material for the experiment) typically have to be optimized for equivalent proteome coverage. For details related to materials and equipment, see the Table of Materials. For details related to buffers and reagent solutions and their composition, see Table 1.

1. Cell surface labeling with biotin

1. Count the cultured cells and pellet approximately 3.5 × 10⁷ live cells by centrifugation (5 min at 500 × g, 24 °C).
   NOTE: The AMO1 plasmacytoma cells were passaged with fresh media every 3 days prior to harvesting.
2. Aspirate the supernatant and resuspend the pellet by adding 1 mL of cold (4 °C) Dulbecco's phosphate-buffered saline (D-PBS) (no calcium, no magnesium), and gently pipetting up and down.
3. Pellet the cells again (as in step 1.1) and repeat the wash step (step 1.2) once more (two washes in total).
4. After the second wash, pellet the cells (as in step 1.1) and resuspend them in 990 µL of cold D-PBS by gently pipetting.
5. Prepare a stock solution of 160 mM sodium metaperiodate (NaIO₄) beforehand. Split into 50 µL aliquots and store at -20 °C for up to 6 months. Add 10 µL of 160 mM sodium metaperiodate solution to the cell suspension in step 1.4, mix thoroughly, and incubate on an end-to-end rotor for 20 min at 4 °C.
6. After the incubation is complete, pellet the cells (as in step 1.1), decant the supernatant, and resuspend in 1 mL of cold D-PBS. Repeat this wash step twice (three washes in total). After the third wash, resuspend the cells in 989 µL of D-PBS.
7. Prepare a stock solution of 100 mM biocytin hydrazide beforehand. Split into 50 µL aliquots, and store at -20 °C for up to 6 months. Add 1 µL of aniline and 10 µL of 100 mM biocytin hydrazide to the cell suspension in step 1.6, mix thoroughly, and incubate on an end-to-end rotor for 60 min at 4 °C.
8. After the incubation is complete, pellet the cells (as in step 1.1), decant the supernatant, and resuspend in 1 mL of cold D-PBS. Repeat this wash step twice (three washes in total).
9. After the third wash, aspirate the supernatant carefully with a pipet, and snap-freeze the cell pellet by placing the tube in liquid N₂. Place the frozen pellet at -80 °C, until ready to proceed with lysis and pulldown.

2. Cell lysis and biotin pulldown

1. Thaw the frozen cell pellet on ice.
2. Add 500 µL of 2x lysis buffer to the pellet, mix thoroughly, and vortex for 30 s.
   NOTE: Vigorous pipetting and vortexing may be required to fully lyse the pellet. Some insoluble debris (i.e., DNA) is acceptable, as it is removed in the subsequent sonication step.
3. Sonicate the cell lysate on ice (three bursts, 20 s each, 60% duty cycle).
4. Centrifuge the lysate to pellet any remaining debris (10 min at 17,200 × g at 4 °C).
5. While the lysate is centrifuging, add 100 µL of neutravidin agarose resin beads to a filtration column. Attach the column to a vacuum manifold, apply a gentle vacuum, and wash the beads by flowing 3 mL of wash buffer 1 over them.
6. Remove the filtration column from the vacuum manifold, cap the bottom, and add the clarified cell lysate to the column with the beads. Cap the top of the column, and incubate the lysate with the beads on an end-to-end rotor for 120 min at 4 °C.
   NOTE: When capping the top of the column, firmly hold the bottom cap in place, otherwise it may get dislodged and the cell lysate may leak during incubation.
7. Prepare wash buffers 1, 2, and 3 (approximately 5 mL of each wash buffer per sample, refer to Table 1), and place them in a 42 °C water bath
8. After the lysate has finished incubation, place the filtration column back onto the vacuum manifold, apply a gentle vacuum, and wash the beads by flowing 5 mL of wash buffer 1 over them.
   NOTE: More than 5 mL of the wash buffers can be used for washing; extra washing may reduce background, non-specific binding onto the beads.
9. Repeat the wash step with 5 mL of wash buffer 2.
10. Repeat the wash step with 5 mL of wash buffer 3.
11. Once the final wash buffer has flowed entirely through the beads, remove the column from the manifold. Add 100 µL of "Lyse" solution to the beads and transfer them to a 1.7 mL low-protein-binding microcentrifuge tube with a pipet. Briefly spin the tube to settle the beads, and remove 50 µL of the solution without disturbing the settled beads.
    NOTE: The reagents in this step and all subsequent steps use a commercially available proteomics sample preparation kit. The kit provides an optimized, simplified sample preparation workflow. However, these steps can also be performed independent of the kit, using a traditional proteomics workflow as described in Verma et al.⁹. If the beads remain stuck to the side or bottom of the column, allow the beads that have been transferred to the tube to settle, and use extra "Lyse" solution in the tube to transfer the remaining beads.
12. Place the tube on a heat block/shaker at 65 °C, and shake at 1,000 rpm for 10 min.
    ​NOTE: This step is required for the reduction and alkylation of cysteine residues prior to digestion.

3. Protein digestion

1. Resuspend the dried trypsin ("Digest" tube in the kit, stored at -20 °C) by adding 210 µL of the "Resuspend" solution. Pipet the solution up and down several times to ensure all the dried trypsin is resuspended.
2. After the incubation is complete, add 50 µL of the resuspended trypsin solution to the beads. Incubate the tube at 37 °C, shaking at 500 rpm for at least 90 min to digest the protein.
3. Once the digestion is complete, transfer the solution containing the beads to a spin column, and insert the column into a 1.7 mL low-protein-binding microcentrifuge tube. Spin the tube (500 × g for 5 min at room temperature [RT]) to separate the digested peptide solution from the beads.
   NOTE: At this stage, PNGase treatment can be performed on the beads once they are separated from the peptide solution, to elute biotinylated peptide fragments from the streptavidin beads. This peptide sample can then be carried forward through the remainder of the workflow as a separate, secondary peptide sample that can be analyzed and compared with the primary sample from on-bead trypsinization. The PNGase approach is likely to provide complementary data to on-bead trypsinization, given the additional specificity for captured N-glycosylated asparagines based on MS-detectable side chain modification¹². However, the drawback of this sequential elution approach is the requirement of twice the mass spectrometer instrument time per sample analysis.
4. Add 100 µL of the "Stop" solution to stop the digestion reaction and acidify the peptide solution.

4. Peptide desalting

1. Using a tube adapter, place a desalting column in a 1.7 mL low-protein-binding microcentrifuge tube. Add the entire acidified peptide solution to the column. Spin the column (3,800 × g for 3 min) so that the solution flows through the column. The peptides are now bound to the column; discard the flowthrough.
2. Add 200 µL of "Wash 1" solution to the column and spin (3,800 × g for 3 min, RT). Discard the flowthrough.
3. Add 200 µL of "Wash 2" solution to the column and spin (3,800 × g for 3 min, RT). Discard the flowthrough.
4. Transfer the column to a new, labeled 1.7 mL low-protein-binding microcentrifuge tube. Add 100 µL of "Elute" solution to the column and spin (3,800 × g for 3 min, RT). Add another 100 µL of "Elute" solution to the column and spin (3,800 × g for 3 min, RT). The peptides are now eluted off the column into the tube.
5. Place the peptide solution in a vacuum centrifuge and allow it to dry overnight. Place the dried-down peptides at -80 °C until ready to quantify, and analyze on the mass spectrometer.

5. Peptide resuspension and quantification

1. Resuspend the dried down peptides in 20 µL of solvent A (0.1% formic acid, 2% acetonitrile).
2. Centrifuge the resuspended peptides (17,200 × g for 10 min) to pellet any insoluble debris.
3. Prepare peptide standards for the colorimetric peptide quantification assay.
   1. Place 5 µL of 1,000 mg/mL peptide stock solution in one well of a 96-well plate.
   2. Perform a seven-step serial dilution in the plate with deionized (DI) water, as follows, with 5 µL of each standard per well: 1,000 mg/mL, 500 mg/mL, 250 mg/mL, 125 mg/mL, 62.5 mg/mL, 31.25 mg/mL, and 15.625 mg/mL. Place 5 µL of DI water in an eighth well for the blank.
4. Place 4 µL of DI water in three separate wells of the 96-well plate. Add 1 µL of the resuspended peptide solution to each well, for a total volume of 5 µL.
   NOTE: When pipetting the peptide solution for quantification, carefully pipet from the top of the solution to prevent disturbing any settled debris.
5. Calculate how much working reagent is needed (45 µL required per well). Prepare the required volume of the colorimetric assay working reagent; vortex the working reagent to ensure proper mixing of the components.
6. Add 45 µL of the working reagent to each of the standard and sample wells.
   NOTE: Make the standards in duplicate, and use a multichannel pipet to ensure a minimal difference in the timing of when the reagent is added to the wells.
7. Cover the plate in aluminum foil and incubate at 37 °C for 15 min.
8. Analyze the plate on an automated plate reader. Use the absorbance values recorded for the standard samples to generate a linear standard curve. Determine the equation of the standard curve and R² value. If the R² value is reasonable (≥0.95), use the standard curve to determine the concentration of resuspended peptides, accounting for the fivefold dilution in the colorimetric assay plate.

6. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis of the digested peptides

1. Adjust the concentration of the peptide samples to 0.2 µg/µL by adding solvent A, and transfer 10 µL into a 0.6 mL low-protein-binding tube.
2. Place the tube in the LC autosampler.
3. Set the LC and MS/MS parameters, according to Figure 2 and Figure 3.
4. Inject 5 µL (1 µg) of the peptide sample into the LC-MS/MS setup for analysis.
   NOTE: The LC-MS/MS settings shown here are only representative for the illustrations. Depending upon the LC and MS instrument availability, users can modify the settings for the LC gradient and MS/MS parameters to obtain deep proteome coverage. For this paper, the MS data analysis was performed using MaxQuant https://www.maxquant.org/, secondary data analysis was performed using Perseus https://maxquant.net/perseus/, and pathway analysis using https://reactome.org/.

Results

For this experiment, we characterized the cell surface proteome of a tumor cell line by labeling N-glycosylated membrane proteins of intact cells with biotin, and enriching these labeled proteins from the whole cell lysate with a neutravidin pulldown (Figure 1). Further, we performed proteome analysis using LC-MS/MS to characterize enriched cell surface proteins. Unlike whole cell proteome analysis, here, the objective was to characterize only cell surface proteins. Hence, we starte...

Discussion

Mass spectrometry-based proteomics is a powerful tool that has enabled unbiased characterization of thousands of unknown proteins on a previously impossible scale. This approach allows us to identify and quantify the proteins, as well as glean a range of insights for the structural and signaling capacities of cells and tissues, by characterizing the variety of proteins present in a particular sample. Moving beyond global protein profiling in a sample, mass spectrometry allows us to characterize various post-translational...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Kits
96X iST Sample Preparation Kit	PreOmics	P.O.00027	Proteomics sample preparation kit. Includes reagents for reduction, alkylation, and digestion. Also include desalting columns and reagents.
Pierce Quantitative Colorimetric Peptide Assay	Thermo	23275	Peptide quantification kit. Includes peptide standards and components of working reagents.
Reagents
Acetonitrile	Fisher	A955-1
Ammonium bicarbonate	Millipore Sigma	09830-1KG
Biocytin hydrazide	Biotium	90060
D-PBS (w/o Calcium and Magnesium Salts)	UCSF Cell Culture Facility	CCFAL003-225B01
Formic Acid	Honeywell	94318
Halt Protease and Phosphatase Inhibitor Single-Use Cocktail	Thermo	1861280
High Capacity Neutravidin Agarose Resin	Thermo	29204
Phosphate Buffered Saline	UCSF Cell Culture Facility	CCFAL001-22J01
RIPA Lysis Buffer, 10x	Millipore Sigma	20-188
Sodium chloride	Fisher	BP358-212
Sodium metaperiodate	Alfa Aesar	13798
Trypan Blue Stain (0.4%)	Gibco	15250-061
Ultrapure 0.5 M EDTA, pH 8.0	Invitrogen	15575-038
Urea (Proteomics Grade)	VWR	M123-1KG
Equipment
TC20 Automated Cell Counter	Bio-Rad	1450102
PrismR Microcentrifuge	Labnet International	C2500-R-230V
Sonicator	VWR	Branson Sonifier 240
Vacuum Manifold	Promega	Promega Vac-Man
Shaking Heatblock	Eppendorf	Eppendorf Thermomixer C
End-to-End rotator	Labnet	Revolver Adjustable Rotator
LC	Thermo	Ultimate 3000 HPLC and UHPLC
Q Exactive Plus Hybrid Quadrapole Orbitrap Mass Spectrometer	Thermo	IQLAAEGAAPFALGMBDK
Microplate Reader	Biotek	Biotek Synergy 2
Vacuum Concentrator	Labconco	7810010
Supplies
1.5 mL Protein LoBind Tubes	Eppendorf	22431081
1.7 mL Microcentrifuge Tubes
Filtration Columns	Bio-Rad	7326008
Spin Columns	Thermo	69725