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.

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
<ol>
	<li>Count the cultured cells and pellet approximately 3.5 &#215; 107 live cells by centrifugation (5 min at 500 &#215; g, 24 &#176;C). 
	NOTE: The AMO1 plasmacytoma cells were passaged with fresh media every 3 days prior to harvesting.</li>
	<li>Aspirate the supernatant and resuspend the pellet by adding 1 mL of cold (4 &#176;C) Dulbecco&#39;s phosphate-buffered saline (D-PBS) (no calcium, no magnesium), and gently pipetting up and down.</li>
	<li>Pellet the cells again (as in step 1.1) and repeat the wash step (step 1.2) once more (two washes in total).</li>
	<li>After the second wash, pellet the cells (as in step 1.1) and resuspend them in 990 &#181;L of cold D-PBS by gently pipetting.</li>
	<li>Prepare a stock solution of 160 mM sodium metaperiodate (NaIO4) beforehand. Split into 50 &#181;L aliquots and store at -20 &#176;C for up to 6 months. Add 10 &#181;L 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 &#176;C.</li>
	<li>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 &#181;L of D-PBS.</li>
	<li>Prepare a stock solution of 100 mM biocytin hydrazide beforehand. Split into 50 &#181;L aliquots, and store at -20 &#176;C for up to 6 months. Add 1 &#181;L of aniline and 10 &#181;L 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 &#176;C.</li>
	<li>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).</li>
	<li>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 &#176;C, until ready to proceed with lysis and pulldown.</li>
</ol>
2. Cell lysis and biotin pulldown
<ol>
	<li>Thaw the frozen cell pellet on ice.</li>
	<li>Add 500 &#181;L 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.</li>
	<li>Sonicate the cell lysate on ice (three bursts, 20 s each, 60% duty cycle).</li>
	<li>Centrifuge the lysate to pellet any remaining debris (10 min at 17,200 &#215; g at 4 &#176;C).</li>
	<li>While the lysate is centrifuging, add 100 &#181;L 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.</li>
	<li>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 &#176;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.</li>
	<li>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 &#176;C water bath</li>
	<li>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.</li>
	<li>Repeat the wash step with 5 mL of wash buffer 2.</li>
	<li>Repeat the wash step with 5 mL of wash buffer 3.</li>
	<li>Once the final wash buffer has flowed entirely through the beads, remove the column from the manifold. Add 100 &#181;L of &#34;Lyse&#34; 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 &#181;L 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&#160;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 &#34;Lyse&#34; solution in the tube to transfer the remaining beads.</li>
	<li>Place the tube on a heat block/shaker at 65 &#176;C, and shake at 1,000 rpm for 10 min. 
	&#8203;NOTE: This step is required for the reduction and alkylation of cysteine residues prior to digestion.</li>
</ol>
3. Protein digestion
<ol>
	<li>Resuspend the dried trypsin (&#34;Digest&#34; tube in the kit, stored at -20 &#176;C) by adding 210 &#181;L of the &#34;Resuspend&#34; solution. Pipet the solution up and down several times to ensure all the dried trypsin is resuspended.</li>
	<li>After the incubation is complete, add 50 &#181;L of the resuspended trypsin solution to the beads. Incubate the tube at 37 &#176;C, shaking at 500 rpm for at least 90 min to digest the protein.</li>
	<li>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 &#215; 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.</li>
	<li>Add 100 &#181;L of the &#34;Stop&#34; solution to stop the digestion reaction and acidify the peptide solution.</li>
</ol>
4. Peptide desalting
<ol>
	<li>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 &#215; g for 3 min) so that the solution flows through the column. The peptides are now bound to the column; discard the flowthrough.</li>
	<li>Add 200 &#181;L of &#34;Wash 1&#34; solution to the column and spin (3,800 &#215; g for 3 min, RT). Discard the flowthrough.</li>
	<li>Add 200 &#181;L of &#34;Wash 2&#34; solution to the column and spin (3,800 &#215; g for 3 min, RT). Discard the flowthrough.</li>
	<li>Transfer the column to a new, labeled 1.7 mL low-protein-binding microcentrifuge tube. Add 100 &#181;L of &#34;Elute&#34; solution to the column and spin (3,800 &#215; g for 3 min, RT). Add another 100 &#181;L of &#34;Elute&#34; solution to the column and spin (3,800 &#215; g for 3 min, RT). The peptides are now eluted off the column into the tube.</li>
	<li>Place the peptide solution in a vacuum centrifuge and allow it to dry overnight. Place the dried-down peptides at -80 &#176;C until ready to quantify, and analyze on the mass spectrometer.</li>
</ol>
5. Peptide resuspension and quantification
<ol>
	<li>Resuspend the dried down peptides in 20 &#181;L of solvent A (0.1% formic acid, 2% acetonitrile).</li>
	<li>Centrifuge the resuspended peptides (17,200 &#215; g for 10 min) to pellet any insoluble debris.</li>
	<li>Prepare peptide standards for the colorimetric peptide quantification assay.
	<ol>
		<li>Place 5 &#181;L of 1,000 mg/mL peptide stock solution in one well of a 96-well plate.</li>
		<li>Perform a seven-step serial dilution in the plate with deionized (DI) water, as follows, with 5 &#181;L 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 &#181;L of DI water in an eighth well for the blank.</li>
	</ol>
	</li>
	<li>Place 4 &#181;L of DI water in three separate wells of the 96-well plate. Add 1 &#181;L of the resuspended peptide solution to each well, for a total volume of 5 &#181;L. 
	NOTE: When pipetting the peptide solution for quantification, carefully pipet from the top of the solution to prevent disturbing any settled debris.</li>
	<li>Calculate how much working reagent is needed (45 &#181;L required per well). Prepare the required volume of the colorimetric assay working reagent; vortex the working reagent to ensure proper mixing of the components.</li>
	<li>Add 45 &#181;L 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.</li>
	<li>Cover the plate in aluminum foil and incubate at 37 &#176;C for 15 min.</li>
	<li>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 (&#8805;0.95), use the standard curve to determine the concentration of resuspended peptides, accounting for the fivefold dilution in the colorimetric assay plate.</li>
</ol>
6. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis of the digested peptides
<ol>
	<li>Adjust the concentration of the peptide samples to 0.2 &#181;g/&#181;L by adding solvent A, and transfer 10 &#181;L into a 0.6 mL low-protein-binding tube.</li>
	<li>Place the tube in the LC autosampler.</li>
	<li>Set the LC and MS/MS parameters, according to Figure 2 and Figure 3.</li>
	<li>Inject 5 &#181;L (1 &#181;g) 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/,&#160;and pathway analysis using https://reactome.org/.</li>
</ol>

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The authors declare no competing financial interests.

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...

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 &#34;cell surface capture&#34;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.

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

"cell surface capture" workflow for label-free quantification of the cell surface proteome

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.

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...

Watch this Scientific Journal Video about "Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome at JoVE.com

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

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.

Over the past decade, mass spectrometry-based proteomics has enabled an in-depth characterization of biological systems across a broad array of ...

cell-surface-capture-workflow-for-label-free-quantification-cell

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

2.1K Views.  University of California, San Francisco. 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.

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

Deep Proteome Profiling by Isobaric Labeling, Extensive Liquid Chromatography, Mass Spectrometry, and Software-assisted Quantification

Many exceptional advances have been made in mass spectrometry (MS)-based proteomics, with particular technical progress in liquid chromatography (LC) coupled to tandem mass spectrometry (LC-MS/MS) and isobaric labeling multiplexing capacity. Here, we introduce a deep-proteomics profiling protocol that combines 10-plex tandem mass tag (TMT) labeling with an extensive LC/LC-MS/MS platform, and post-MS computational interference correction to accurately quantitate whole proteomes. This protocol includes the following main steps: protein extraction and digestion, TMT labeling, 2-dimensional (2D) LC, high-resolution mass spectrometry, and computational data processing. Quality control steps are included for troubleshooting and evaluating experimental variation. More than 10,000 proteins in mammalian samples can be confidently quantitated with this protocol. This protocol can also be applied to the quantitation of post translational modifications with minor changes. This multiplexed, robust method provides a powerful tool for proteomic analysis in a variety of complex samples, including cell culture, animal tissues, and human clinical specimens.

We present a protocol to accurately quantitate proteins with isobaric labelling, extensive fractionation, bioinformatics tools, and quality control steps in combination with liquid chromatography interfaced to a high-resolution mass spectrometer.

Many exceptional advances have been made in mass spectrometry (MS)-based proteomics, with particular technical progress in liquid chromatography (LC) ...

Quantitative Analysis of Chromatin Proteomes in Disease

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due to the high percentage of binucleated cells,1 the predominantly heterochromatic state of the DNA, and the non-dividing nature of the cardiomyocyte which renders adult nuclei in a permanent state of interphase.2 Transcriptional regulation during development and disease have been well studied in this organ,3-5 but what remains relatively unexplored is the role played by the nuclear proteins responsible for DNA packaging and expression, and how these proteins control changes in transcriptional programs that occur during disease.6 In the developed world, heart disease is the number one cause of mortality for both men and women.7 Insight on how nuclear proteins cooperate to regulate the progression of this disease is critical for advancing the current treatment options.
Mass spectrometry is the ideal tool for addressing these questions as it allows for an unbiased annotation of the nuclear proteome and relative quantification for how the abundance of these proteins changes with disease. While there have been several proteomic studies for mammalian nuclear protein complexes,8-13 until recently14 there has been only one study examining the cardiac nuclear proteome, and it considered the entire nucleus, rather than exploring the proteome at the level of nuclear sub compartments.15 In large part, this shortage of work is due to the difficulty of isolating cardiac nuclei. Cardiac nuclei occur within a rigid and dense actin-myosin apparatus to which they are connected via multiple extensions from the endoplasmic reticulum, to the extent that myocyte contraction alters their overall shape.16 Additionally, cardiomyocytes are 40% mitochondria by volume17 which necessitates enrichment of the nucleus apart from the other organelles. Here we describe a protocol for cardiac nuclear enrichment and further fractionation into biologically-relevant compartments. Furthermore, we detail methods for label-free quantitative mass spectrometric dissection of these fractions-techniques amenable to in vivo experimentation in various animal models and organ systems where metabolic labeling is not feasible.

Advances in mass spectrometry have allowed the high throughput analysis of protein expression and modification in a host of tissues. Combined with subcellular fractionation and disease models, quantitative mass spectrometry and bioinformatics can reveal new properties in biological systems. The method described herein analyzes chromatin-associated proteins in the setting of heart disease and is readily applicable to other in vivo models of human disease.

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due ...

Medicine

Selective Labelling of Cell-surface Proteins using CyDye DIGE Fluor Minimal Dyes

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of almost all intracellular signaling. Moreover, they play an important role in environmental adaptation and drug treatment, and are often involved in disease pathogenesis and pathology (1). Protein-protein interactions are intrinsic to signaling pathways, and to gain more insight in these complex biological processes, sensitive and reliable methods are needed for studying cell surface proteins. Two-dimensional (2-D) electrophoresis is used extensively for detection of biomarkers and other targets in complex protein samples to study differential changes. Cell surface proteins, partly due to their low abundance (1 2% of cellular proteins), are difficult to detect in a 2-D gel without fractionation or some other type of enrichment. They are also often poorly represented in 2-D gels due to their hydrophobic nature and high molecular weight (2). In this study, we present a new protocol for intact cells using CyDye  DIGE Fluor minimal dyes for specific labeling and detection of this important group of proteins. The results showed specific labeling of a large number of cell surface proteins with minimal labeling of intracellular proteins. This protocol is rapid, simple to use, and all three CyDye DIGE Fluor minimal dyes (Cy 2, Cy 3 and Cy 5) can be used to label cell-surface proteins. These features allow for multiplexing using the 2-D Fluorescence Difference Gel Electrophoresis (2-D DIGE) with Ettan  DIGE technology and analysis of protein expression changes using DeCyder  2-D Differential Analysis Software. The level of cell-surface proteins was followed during serum starvation of CHO cells for various lengths of time (see Table 1). Small changes in abundance were detected with high accuracy, and results are supported by defined statistical methods.

A simple and specific method was demonstrated for fluorescent labeling and enhanced detection of cell surface proteins without a fractionation step. Differential abundance in cell surface proteins was analyzed using two-dimensional (2-D) electrophoresis and Ettan&#x2122; DIGE technology.

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of ...

Biology

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

Label-Free Immunoprecipitation Mass Spectrometry Workflow for Large-scale Nuclear Interactome Profiling

Immunoaffinity purification mass spectrometry (IP-MS) has emerged as a robust quantitative method of identifying protein-protein interactions. This publication presents a complete interaction proteomics workflow designed for identifying low abundance protein-protein interactions from the nucleus that could also be applied to other subcellular compartments. This workflow includes subcellular fractionation, immunoprecipitation, sample preparation, offline cleanup, single-shot label-free mass spectrometry, and downstream computational analysis and data visualization. Our protocol is optimized for detecting compartmentalized, low abundance interactions that are difficult to identify from whole cell lysates (e.g., transcription factor interactions in the nucleus) by immunoprecipitation of endogenous proteins from fractionated subcellular compartments. The sample preparation pipeline outlined here provides detailed instructions for the preparation of HeLa cell nuclear extract, immunoaffinity purification of endogenous bait protein, and quantitative mass spectrometry analysis. We also discuss methodological considerations for performing large-scale immunoprecipitation in mass spectrometry-based interaction profiling experiments and provide guidelines for evaluating data quality to distinguish true positive protein interactions from nonspecific interactions. This approach is demonstrated here by investigating the nuclear interactome of the CMGC kinase, DYRK1A, a low abundance protein kinase with poorly defined interactions within the nucleus.

Described is a proteomics workflow for identifying protein interaction partners from a nuclear subcellular fraction using immunoaffinity enrichment of a given protein of interest and label-free mass spectrometry. The workflow includes subcellular fractionation, immunoprecipitation, filter aided sample preparation, offline cleanup, mass spectrometry, and a downstream bioinformatics pipeline.

Immunoaffinity purification mass spectrometry (IP-MS) has emerged as a robust quantitative method of identifying protein-protein interactions. This ...

Label-Free Imaging of Single Proteins Secreted from Living Cells via iSCAT Microscopy

We demonstrate interferometric scattering (iSCAT) microscopy, a method capable of detecting single unlabeled proteins secreted from individual living cells in real time. In this protocol, we cover the fundamental steps to realize an iSCAT microscope and complement it with additional imaging channels to monitor the viability of a cell under study. Following this, we use the method for real-time detection of single proteins as they are secreted from a living cell which we demonstrate with an immortalized B-cell line (Laz388). Necessary steps concerning the preparation of microscope and sample as well as the analysis of the recorded data are discussed. The video protocol demonstrates that iSCAT microscopy offers a straightforward method to study secretion at the single-molecule level.

We present a protocol for the real-time optical detection of single unlabeled proteins as they are secreted from living cells. This is based on interferometric scattering (iSCAT) microscopy, which can be applied to a variety of different biological systems and configurations.

We demonstrate interferometric scattering (iSCAT) microscopy, a method capable of detecting single unlabeled proteins secreted from individual living ...

Engineering

Label-Free Quantitative Proteomics Workflow for Discovery-Driven Host-Pathogen Interactions

The technological achievements of mass spectrometry (MS)-based quantitative proteomics opens many undiscovered avenues for analyzing an organism&#8217;s global proteome under varying conditions. This powerful strategy applied to the interactions of microbial pathogens with the desired host comprehensively characterizes both perspectives towards infection. Herein, the workflow describes label-free quantification (LFQ) of the infectome of Cryptococcus neoformans, a fungal facultative intracellular pathogen that is the causative agent of the deadly disease cryptococcosis, in the presence of immortalized macrophage cells. The protocol details the proper protein preparation techniques for both pathogen and mammalian cells within a single experiment, resulting in appropriate peptide submission for liquid-chromatography (LC)-MS/MS analysis. The high throughput generic nature of LFQ allows a wide dynamic range of protein identification and quantification, as well as transferability to any host-pathogen infection setting, maintaining extreme sensitivity. The method is optimized to catalogue extensive, unbiased protein abundance profiles of a pathogen within infection-mimicking conditions. Specifically, the method demonstrated here provides essential information on C. neoformans pathogenesis, such as protein production necessary for virulence and identifies critical host proteins responding to microbial invasion.

Here, we present a protocol to profile the interplay between host and pathogen during infection by mass spectrometry-based proteomics. This protocol uses label-free quantification to measure changes in protein abundance of both host (e.g., macrophages) and pathogen (e.g., Cryptococcus neoformans) in a single experiment.

The technological achievements of mass spectrometry (MS)-based quantitative proteomics opens many undiscovered avenues for analyzing an organism&#8217;s ...

Immunology and Infection

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...

Workflow for High-content, Individual Cell Quantification of Fluorescent Markers from Universal Microscope Data, Supported by Open Source Software

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly dependent on modes of single-cell analysis. Methods which deliver composite data reflecting the mean values of biomarkers from cell populations risk losing subpopulation dynamics that reflect the heterogeneity of the studied biological system. In keeping with this, traditional approaches are being replaced by, or supported with, more sophisticated forms of cellular assay developed to allow assessment by high-content microscopy. These assays potentially generate large numbers of images of fluorescent biomarkers, which enabled by accompanying proprietary software packages, allows for multi-parametric measurements per cell. However, the relatively high capital costs and overspecialization of many of these devices have prevented their accessibility to many investigators.
Described here is a universally applicable workflow for the quantification of multiple fluorescent marker intensities from specific subcellular regions of individual cells suitable for use with images from most fluorescent microscopes. Key to this workflow is the implementation of the freely available Cell Profiler software1&#160;to distinguish individual cells in these images, segment them into defined subcellular regions and deliver fluorescence marker intensity values specific to these regions. The extraction of individual cell intensity values from image data is the central purpose of this workflow and will be illustrated with the analysis of control data from a siRNA screen for G1 checkpoint regulators in adherent human cells. However, the workflow presented here can be applied to analysis of data from other means of cell perturbation (e.g., compound screens) and other forms of fluorescence based cellular markers and thus should be useful for a wide range of laboratories.

Presented is a flexible informatics workflow enabling multiplexed image-based analysis of fluorescently labeled cells. The workflow quantifies nuclear and cytoplasmic markers and computes marker translocation between these compartments. Procedures are provided for perturbation of cells using siRNA and reliable methodology for marker detection by indirect immunofluorescence in 96-well formats.

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly ...

High-throughput and Deep-proteome Profiling by 16-plex Tandem Mass Tag Labeling Coupled with Two-dimensional Chromatography and Mass Spectrometry

Isobaric tandem mass tag (TMT) labeling is widely used in proteomics because of its high multiplexing capacity and deep proteome coverage. Recently, an expanded 16-plex TMT method has been introduced, which further increases the throughput of proteomic studies. In this manuscript, we present an optimized protocol for 16-plex TMT-based deep-proteome profiling, including protein sample preparation, enzymatic digestion, TMT labeling reaction, two-dimensional reverse-phase liquid chromatography (LC/LC) fractionation, tandem mass spectrometry (MS/MS), and computational data processing. The crucial quality control steps and improvements in the process specific for the 16-plex TMT analysis are highlighted. This multiplexed process offers a powerful tool for profiling a variety of complex samples such as cells, tissues, and clinical specimens. More than 10,000 proteins and posttranslational modifications such as phosphorylation, methylation, acetylation, and ubiquitination in highly complex biological samples from up to 16 different samples can be quantified in a single experiment, providing a potent tool for basic and clinical research.

Presented here is an optimized high-throughput protocol developed with 16-plex tandem mass tag reagents, enabling quantitative proteome profiling of biological samples. Extensive basic pH fractionation and high-resolution LC-MS/MS mitigate ratio compression and provide deep proteome coverage.

Isobaric tandem mass tag (TMT) labeling is widely used in proteomics because of its high multiplexing capacity and deep proteome coverage. Recently, an ...

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

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Przeglądaj więcej filmów

Deep Proteome Profiling by Isobaric Labeling, Extensive Liquid Chromatography, Mass Spectrometry, and Software-assisted Quantification

Quantitative Analysis of Chromatin Proteomes in Disease

Selective Labelling of Cell-surface Proteins using CyDye DIGE Fluor Minimal Dyes

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Label-Free Immunoprecipitation Mass Spectrometry Workflow for Large-scale Nuclear Interactome Profiling

Label-Free Imaging of Single Proteins Secreted from Living Cells via iSCAT Microscopy

Label-Free Quantitative Proteomics Workflow for Discovery-Driven Host-Pathogen Interactions

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Workflow for High-content, Individual Cell Quantification of Fluorescent Markers from Universal Microscope Data, Supported by Open Source Software

High-throughput and Deep-proteome Profiling by 16-plex Tandem Mass Tag Labeling Coupled with Two-dimensional Chromatography and Mass Spectrometry