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.

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		</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.&#160;</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.&#160;</td>
		</tr>
		<tr>
			<td>Reagents</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>Equipment</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&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum Concentrator</td>
			<td>Labconco</td>
			<td>7810010</td>
			<td></td>
		</tr>
		<tr>
			<td>Supplies</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

Research

JoVE Journal

Biochemistry

1.9K 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

Purification of Biotinylated Cell Surface Proteins from Rhipicephalus microplus Epithelial Gut Cells

Rhipicephalus microplus &#8211; the cattle tick &#8211; is the most significant ectoparasite in terms of economic impact on livestock as a vector of several pathogens. Efforts have been dedicated to the cattle tick control to diminish its deleterious effects, with focus on the discovery of vaccine candidates, such as BM86, located on the surface of the tick gut epithelial cells. Current research focuses upon the utilization of cDNA and genomic libraries, to screen for other vaccine candidates. The isolation of tick gut cells constitutes an important advantage in investigating the composition of surface proteins upon the tick gut cells membrane. This paper constitutes a novel and feasible method for the isolation of epithelial cells, from the tick gut contents of semi-engorged R. microplus. This protocol utilizes TCEP and EDTA to release the epithelial cells from the subepithelial support tissues and a discontinuous density centrifugation gradient to separate epithelial cells from other cell types. Cell surface proteins were biotinylated and isolated from the tick gut epithelial cells, using streptavidin-linked magnetic beads allowing for downstream applications in FACS or LC-MS/MS-analysis.

Purification of Biotinylated Cell Surface Proteins from Rhipicephalus microplus Epithelial Gut Cells

A modified density centrifugation gradient-based methodology was utilized to isolate epithelial cells from Rhipicephalus microplus gut tissue. Surface-bound proteins were biotinylated and purified through streptavidin magnetic beads for utilization in downstream applications.

Rhipicephalus microplus &#8211; the cattle tick &#8211; is the most significant ectoparasite in terms of economic impact on livestock as a vector of ...

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) ...

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and podocytes with their slit diaphragms. The delicate structure of the glomerular filter, especially the slit diaphragm, relies on the interplay of diverse cell surface proteins. Studying these cell surface proteins has so far been limited to in vitro studies or histologic analysis. Here, we present a murine in vivo biotinylation labeling method, which enables the study of glomerular cell surface proteins under physiologic and pathophysiologic conditions. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of a protein of interest. Semi-quantitation of glomerular cell surface abundance is readily available with this novel method, and all proteins accessible to biotin perfusion and immunoprecipitation can be studied. In addition, isolation of glomeruli with or without biotinylation enables further analysis of glomerular RNA and protein as well as primary glomerular cell culture (i.e., primary podocyte cell culture).

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Here we present a protocol for murine in vivo labeling of glomerular cell surface proteins with biotin. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of the protein of interest.

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and ...

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 ...

Real-Time, Semi-Automated Fluorescent Measurement of the Airway Surface Liquid pH of Primary Human Airway Epithelial Cells

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) hydration, mucus viscosity and activity of antimicrobial peptides, key parameters involved in innate defense of the lungs. This is of primary relevance in the field of chronic respiratory diseases such as cystic fibrosis (CF) where these parameters are dysregulated. While different groups have studied ASL pH both in vivo and in vitro, their methods report a relatively wide range of ASL pH values and even contradictory findings regarding any pH differences between non-CF and CF cells. Furthermore, their protocols do not always provide enough details in order to ensure reproducibility, most are low throughput and require expensive equipment or specialized knowledge to implement, making them difficult to establish in most labs. Here we describe a semi-automated fluorescent plate reader assay that enables the real-time measurement of ASL pH under thin film conditions that more closely resemble the in vivo situation. This technique allows for stable measurements for many hours from multiple airway cultures simultaneously and, importantly, dynamic changes in ASL pH in response to agonists and inhibitors can be monitored. To achieve this, the ASL of fully differentiated primary human airway epithelial cells (hAECs) are stained overnight with a pH-sensitive dye in order to allow for the reabsorption of the excess fluid to ensure thin film conditions. After fluorescence is monitored in the presence or absence of agonists, pH calibration is performed in situ to correct for volume and dye concentration. The method described provides the required controls to make stable and reproducible ASL pH measurements, which ultimately could be used as a drug discovery platform for personalized medicine, as well as adapted to other epithelial tissues and experimental conditions, such as inflammatory and/or host-pathogen models.

We present a protocol to make dynamic measurements of the airway surface liquid pH under thin film conditions using a plate-reader.

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) ...

Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the structure-function relationship of these receptors in situ, we need to visualize and track them on the live cell surface with enough spatiotemporal resolution. Here we show how to use recently developed Lattice Light-Sheet Microscopy (LLSM) to image T-cell receptors (TCRs) four-dimensionally (4D, space and time) at the live cell membrane. T cells are one of the main effector cells of the adaptive immune system, and here we used T cells as an example to show that the signaling and function of these cells are driven by the dynamics and interactions of the TCRs. LLSM allows for 4D imaging with unprecedented spatiotemporal resolution. This microscopy technique therefore can be generally applied to a wide array of surface or intracellular molecules of different cells in biology.

The goal of this protocol is to show how to use Lattice Light-Sheet Microscopy to four-dimensionally visualize surface receptor dynamics in live cells. Here T cell receptors on CD4+ primary T cells are shown.

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the ...

SUMO-Binding Entities (SUBEs) as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in liver cancer, the modification by SUMO (Small Ubiquitin MOdifier) has been given the most attention. Isolation of endogenous SUMOylated proteins in vivo is challenging due to the presence of active SUMO-specific proteases. Initial studies of SUMOylation in vivo were based on the molecular detection of specific SUMOylated proteins (e.g., by western blot). However, in many cases, antibodies, generally made with non-modified recombinant protein, did not immunoprecipitate SUMOylated forms of the protein of interest.&#160;Nickel chromatography has been the other approach to study SUMOylation by capturing histidine-tagged versions of SUMO molecules. This approach is mainly used in cells stably expressing or transiently transfected with His-SUMO molecules. To overcome these limitations, SUMO-binding entities (SUBEs) were developed to isolate endogenous SUMOylated proteins. Herein, we describe all the steps required for the enrichment, isolation, and identification of SUMOylated substrates from human hepatoma cells and hepatic tissues from a liver cancer mouse model by using SUBEs. Firstly, we describe methods involved in the preparation and lysis of the human hepatoma cells and liver tumor tissue samples. Then, a thorough explanation of the preparation of SUBEs and controls is detailed along with the protocol for the protein pull-down assays. Finally, some examples are provided regarding the options available for the identification and characterization of the SUMOylated proteome, namely the use of western-blot analysis for the detection of a specific SUMOylated substrate from liver tumors or the use of proteomics by mass spectrometry for high-throughput characterization of the SUMOylated proteome and interactome in hepatoma cells.

SUMO-Binding Entities SUBEs as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Here, we present a protocol to enrich, isolate, identify, and characterize proteins modified by SUMO in vivo both from human hepatoma cells and liver tumors obtained from mouse models of hepatocellular carcinoma by using SUMO-binding entities (SUBEs).

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in ...

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 ...

Stepwise Dosing Protocol for Increased Throughput in Label-Free Impedance-Based GPCR Assays

Label-free impedance-based assays are increasingly used to non-invasively study ligand-induced GPCR activation in cell culture experiments. The approach provides real-time cell monitoring with a device-dependent time resolution down to several tens of milliseconds and it is highly automated. However, when sample numbers get high (e.g., dose-response studies for various different ligands), the cost for the disposable electrode arrays as well as the available time resolution for sequential well-by-well recordings may become limiting. Therefore, we here present a serial agonist addition protocol which has the potential to significantly increase the output of label-free GPCR assays. Using the serial agonist addition protocol, a GPCR agonist is added sequentially in increasing concentrations to a single cell layer while continuously monitoring the sample&#39;s impedance (agonist mode). With this serial approach, it is now possible to establish a full dose-response curve for a GPCR agonist from just one single cell layer. The serial agonist addition protocol is applicable to different GPCR coupling types, Gq Gi/0 or Gs and it is compatible with recombinant and endogenous expression levels of the receptor under study. Receptor blocking by GPCR antagonists is assessable as well (antagonist mode).

This protocol demonstrates real-time recording of full dose-response relationships for agonist-induced GPCR activation from a single cell layer grown on a single microelectrode using label-free impedance measurements. The new dosing scheme significantly increases throughput without loss in time resolution.

Label-free impedance-based assays are increasingly used to non-invasively study ligand-induced GPCR activation in cell culture experiments. The approach ...

RNA Blot Analysis for the Detection and Quantification of Plant MicroRNAs

MicroRNAs (miRNAs) are a class of endogenously expressed non-coding, ~21 nt small RNAs involved in the regulation of gene expression in both plants and animals. Most miRNAs act as negative switches of gene expression targeting key genes. In plants, primary miRNAs (pri-miRNAs) transcripts are generated by RNA polymerase II, and they form varying lengths of stable stem-loop structures called pre-miRNAs. An endonuclease, Dicer-like1, processes the pre-miRNAs into miRNA-miRNA* duplexes. One of the strands from miRNA-miRNA* duplex is selected and loaded onto Argonaute 1 protein or its homologs to mediate the cleavage of target mRNAs. Although miRNAs are key signaling molecules, their detection is often carried out by less than optimal PCR-based methods instead of a sensitive northern blot analysis. We describe a simple, reliable, and extremely sensitive northern method that is ideal for the quantification of miRNA levels with very high sensitivity, literally from any plant tissue. Additionally, this method can be used to confirm the size, stability and the abundance of miRNAs and their precursors.

This method demonstrates use of the northern hybridization technique to detect miRNAs from total RNA extract.

MicroRNAs (miRNAs) are a class of endogenously expressed non-coding, ~21 nt small RNAs involved in the regulation of gene expression in both plants and ...