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JoVE Journal Biochemistry

Biochemistry

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

Published: March 24, 2023 doi: 10.3791/64952
Automatic Translation
1, 1,2, 1,3,4
1Department of Laboratory Medicine, University of California, San Francisco, 2Department of Biosciences and Bioengineering, Indian Institute of Technology Bombay, 3Deptartment of Bioengineering and Therapeutic Sciences, University of California, San Francisco, 4Chan Zuckerberg Biohub

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.

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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/.

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Representative 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 started with a sample input of 3.5 × 107 AMO-1 plasmacytoma cells. We obtained a final peptide concentration of ~630 ng/µL based on the colorimetric peptide quantification assay, and the total yield was ~12.6 µg of peptides (in 20 µL) (Figure 2A). We then injected 1 µg of peptide sample into the LC-MS/MS system, and repeated the injection three times for technical replicates. The LC and MS parameters used were optimized based on prior experiments (Figure 2C and Figure 3A). We utilized a long LC gradient of 4 h to ensure maximum coverage.

LC-MS/MS data were analyzed using MaxQuant; we were able to identify 2,221 proteins with at least one unique peptide, and 1,764 proteins with at least two unique peptides across all the three replicates. In total, there were 12,307 unique peptides identified. We determined cell surface protein enrichment in the sample by utilizing a Python script (the exact script we used can be found in Supplementary Figure S1) that compares the results obtained from MaxQuant to several established datasets of cell surface proteins. These datasets include the previously reported "in silico surfaceome"3 and a dataset established from proteins annotated as membrane-localized in UniProt13 (datasets are available in Supplementary Table S1). This analysis yielded 601 surface proteins in our sample (approximately 27% of all the proteins identified) (Figure 4A). Gene ontology annotation (http://geneontology.org/) was performed, which indicated approximately 30% of proteins to be canonically localized to the plasma membrane. Reactome pathway analysis (https://reactome.org/) also indicated relative enrichment of the membrane proteins involved in trans-membrane transport and cell surface interactions (Figure 5 and Supplementary Table S2).

Figure 1
Figure 1: Workflow for cell surface labeling and subsequent sample preparation for mass spectrometry analysis. First, proteins at the surface of the desired cellular sample are labeled with biotin. These cells are then lysed, followed by incubation with neutravidin beads that bind biotin-labeled proteins. The beads are then washed repeatedly to remove intracellular proteins and other contaminants. The protein sample then undergoes an on-bead digestion with the addition of trypsin. The resulting sample of peptide fragments then undergoes desalting to remove contaminants from the washing and digestion steps. The final peptide sample is then dried down and ready to undergo mass spectrometry analysis. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Colorimetric peptide quantification standard curve and LC gradient. (A) Eight-point standard curve generated by colorimetric peptide quantification assay used to determine peptide concentration in processed samples. (B) An illustration of the LC-MS/MS system used to process samples. (C) Liquid chromatography gradient and flow rate used for injection of samples onto the mass spectrometer. Abbreviations: LC-MS/MS = liquid chromatography-tandem mass spectrometry. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Mass spectrometry parameters and representative chromatogram. (A) Parameters used for this experiment using an Orbitrap mass spectrometer. (B) Representative chromatogram generated by mass spectrometry analysis following "cell surface capture" protocol for a 4 h gradient. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Cell surface protein enrichment analysis and comparison of technical replicates. These datasets include the previously reported "in silico surfaceome"3 and a dataset established from proteins annotated as membrane-localized in UniProt13. (A) The number of proteins and peptides identified with a different cut-off after removing contaminants, reverse proteins, and a q-value > 0.05. (B) The plot represents the distribution of the identified protein and Andromeda score14. (C) The scatter plot illustrates the sample-to-sample correlation. The Spearman's rank correlation measures the strength and direction of association between two ranked variables. (D) Box plots depicting the run-to-run variation. (E) Sample-wise distribution plot representing the data quality of replicates. Abbreviation: LFQ = label-free quantitation. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Pathway analysis. Pathway analysis was performed using Reactome version 8222. An illustrative depiction of cell surface interactions at the vascular wall, exhibiting the enrichment of cell surface proteome for key interactions involved in the platelet and leukocyte interaction with the endothelium. Please click here to view a larger version of this figure.

Reagent Name Composition Storage
100 mM biocytin hydrazide  powder form biocytin hydrazide dissolved in DMSO Split into 50 μL aliquots and store at -20 °C for up to 6 months. 
160 mM sodium metaperiodate (NaIO4) solid NaIO4 dissolved in DI water Split into 50 μL aliquots and store at -20 °C for up to 6 months. 
2x RIPA Lysis Buffer 2x RIPA Lysis Buffer, 1.25 mM EDTA, Single Use Protease Inhibitor Cocktail Prepare fresh each time before using 10x RIPA Lysis Buffer.
Peptide Colorimetric Assay Working Reagent 50% Reagent A, 48% Reagent B, 2% Reagent C Prepare fresh each time before use. Store components at 4 °C.
Solvent A 0.1% Formic Acid, 2% Acetonitrile Can prepare ahead of time and store at room temperature
Wash Buffer 1 1x RIPA Lysis Buffer, 1 mM EDTA Can prepare large batch beforehand and store at room temperature
Wash Buffer 2 1x PBS, 1 M NaCl Can prepare large batch beforehand and store at room temperature
Wash Buffer 3 2 M Urea, 50 mM Ammonium bicarbonate Prepare fresh each time as urea will degrade quickly over time.

Table 1: Buffers and reagent solutions used in this protocol.

Supplementary Figure S1: Python script for comparing results obtained from MaxQuant to several established datasets of cell surface proteins. Please click here to download this File.

Supplementary Table S1: Datasets of cell surface proteins. These datasets include the previously reported "in silico surfaceome"3 and a dataset established from proteins annotated as membrane-localized in UniProt13. Please click here to download this File.

Supplementary Table S2: The 10 most relevant pathways (sorted by p value) are presented here. Abbreviations: FDR = false discovery rate; SLC = solute carrier; TCR = T cell receptor. Please click here to download this File.

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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 modifications (PTMs) on these proteins, providing an even deeper glimpse into the stages of signaling pathways and protein dynamics not available when analyzing at the DNA or RNA level1. Toward the development of novel cancer therapeutics, mass spectrometry-based proteomics allows us to compare the protein profile of malignant tumor cells versus healthy tissues, to identify potential druggable targets.

The rapidly growing field of cancer immunotherapy relies on antigens expressed on the surface of malignant cells to identify and selectively destroy tumors. Since the development of novel cancer immunotherapies is limited by the lack of unique cancer-specific antigens, which are absent in healthy tissues, it is critical to identify more novel antigens specific to tumor cells15. Tumor specific antigens need not be whole proteins unique to the tumor cell, but can also be specific protein conformations or PTMs absent in healthy tissues16,17. Proteomic analysis of cell surface proteins on both tumor and healthy cells can yield novel cancer-specific antigen targets. Cell surface proteomics requires the enrichment of surface proteins from the whole-cell lysate prior to mass spectrometry analysis, to reduce the complexity of the sample and avoid ion suppression of the desired membrane proteins (often low abundance) by highly expressed intracellular proteins18. While the cell surface capture strategy employed here is one of the most commonly used enrichment approaches for cell surface proteomics, there are other methods for cell surface protein enrichment that have been integrated with a mass spectrometry workflow, including selective ultracentrifugation of cellular membranes, NHS-biotin labeling of surface lysines, and enzyme-mediated biotinylation of cell surface proteins19,20. Head-to-head comparisons suggest that the cell surface capture approach provides the optimal balance of ease-of-use and successful, relatively unbiased enrichment of cell surface proteins with the lowest background labeling of intracellular proteins20.

This paper presents an effective and time efficient protocol for a cell surface proteomics workflow by integrating the cell surface labeling and pulldown procedure with an optimized proteomics sample preparation workflow (Figure 1). The entire procedure, from harvesting cultured cells to analysis on the mass spectrometer, could be achieved over the course of 3 days. To obtain most robust coverage of the surface proteome, the initial input of cells may need to be optimized based on the cell type being used. Performing a pilot experiment with varying cell input, ranging from a few million cells up to 50 million cells, is recommended to ensure maximal coverage. Additionally, if a large amount of highly expressing intracellular proteins is observed, increased washing of the neutravidin beads following incubation with the lysate could minimize non-specific binding of proteins to the beads.

We recommend performing the experiment with at least three biological replicates and three technical replicates per condition, to enhance confidence that the identified cell surface markers are indeed present on the majority of cells. Additionally, when performing the cell surface capture workflow for a new sample type, it could be helpful to analyze a whole-cell lysate using a conventional shotgun proteomics approach21, to compare with the cell surface capture results. Analysis of the raw mass spectrometry data can be performed by various database search and software packages-here, we used MaxQuant-to yield a list of identified proteins22,23. This list can be filtered against various known databases3,24 of surface proteins to establish a list of the cell surface proteins identified in a single experiment. Once a list of cell surface proteins has been established, they can be further analyzed for their role in relevant biochemical pathways25 or drug target candidates26. This workflow can be applied to a wide variety of cell types, including suspension and adherent cell lines (for adherent cells, we recommend lifting cells without the use of trypsin prior to sample preparation, to avoid cleavage of the cell surface proteins), as well as primary samples.

Using this workflow, we were able to confidently characterize the cell surface proteome of a particular cell line. By comparing the surface proteome of tumor cell lines against healthy cells, and cross-referencing with an RNA sequencing database for normal tissues, a list of potential immunotherapeutic targets could be established8,13. A limitation of this method is that, despite the significant enrichment of cell surface proteins versus a standard whole-cell lysate experiment, the majority of the proteins identified by mass spectrometry analysis in this workflow are still annotated as intracellular. Depending on the stringency of the surface protein database used, approximately 25%-30% of the proteins identified are from the cell surface. We anticipate that a significant part of the non-surface background likely emanates from background protein non-specifically bound to beads (for example, as characterized by CRAPome27 analysis), while others represent cell penetration of the labeling reagent to react with N-glycosylated proteins in the endoplasmic reticulum, Golgi, or other intracellular compartments.

While alternate peptide elution methods such as PNGase treatment result in much greater enrichment of N-glycosites among the total peptides analyzed, this approach only allows the identification of individual glycyosylated peptides, and loses valuable information from the non-glycosylated remainder of the pulled-down protein sequence12. This extra information can be captured with this on-bead trypsinization approach, though at the potential tradeoff of reduced specificity for N-glycosylated proteins analyzed in the mass spectrometer. In addition, even PNGase elution does not overcome the challenge of labeling intracellular glycosylated proteins, and a PNGase elution leads to typically only a single peptide per protein; this leads to significant variability in protein identification, per our lab's experience, whereas the on-bead digestion described here leads to multiple peptides per captured protein. Therefore, depending on the objective of the experiment, one could potentially choose different strategies for employing the cell surface capture strategy. These strategies include miniaturization and automation of the cell surface capture protocol for small sample inputs28 and utilizing streptavidin beads of varying binding capacity29 depending on the application. The method and workflow illustrated here serves as a robust, easy-to-perform general strategy for cell surface protein enrichment.

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Disclosures

The authors declare no competing financial interests.

Acknowledgments

We thank Dr. Kamal Mandal (Dept of Laboratory Medicine, UCSF) for help with setting up the LC-MS/MS run, Deeptarup Biswas (BSBE, IIT Bombay) for help with data analysis, and Dr. Audrey Reeves (Dept of Laboratory Medicine, UCSF) for help with data analysis. Related work in the A.P.W. lab is supported by NIH R01 CA226851 and the Chan Zuckerberg Biohub. Figure 1 and Figure 2B were made using BioRender.com.

Materials

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

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References

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Tags

Cell Surface Capture Label-free Quantification Cell Surface Proteome Antigens Cell Membrane Proteins Mass Spectrometry-based Proteomics Tumor Cells Cancer Immunotherapies Biotin-labeled Cell Pellet Lysis Buffer Sonication Centrifugation NeutrAvidin Agarose Resin Beads Filtration Column
"Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome
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Naik, A., Srivastava, S., Wiita, A.… More

Naik, A., Srivastava, S., Wiita, A. P. "Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome. J. Vis. Exp. (193), e64952, doi:10.3791/64952 (2023).

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