This research has been supported by the startup fund of Simon Fraser University. &#160;

1. Harvest Membranes
<ol>
	<li>Add hypotonic buffer (10 mM Tris-HCl, 10 mM KCl, 1.5 mM MgCl2, pH 8.0) with protease inhibitor cocktail onto a cell pellet (~108 cells) and incubate for 15-30 min on ice. Lyse the cells by passing the sample through syringe needles (5-10 passes) or homogenizing the sample by a Dounce homogenizer (15-30 strokes). Use a hemocytometer and trypan blue staining to check the efficiency of lysis.</li>
	<li>Obtain the microsomal fraction by a differential centrifugation of the lysate at 3,000 g x 15 min and then at 100,000 g x 2 hr. The first centrifugation obtains a pellet that is the nuclear fraction, and the subsequent centrifugation of the supernatant generates a second pellet, which is the microsomal fraction that contains the plasma membrane as well as membrane-bound organelles8. The final supernatant is the cytosolic fraction. Store the microsomal fractions in -80 &#176;C freezer for further analysis as indicated below.</li>
</ol>
2. Dissolve, Denature, and Digest Membrane Proteins
<ol>
	<li>Dissolve the microsomal fraction in the denaturation buffer (40 mM Tris, 10 mM EDTA, 10 mM TCEP, 0.5% Rapigest, pH 8) and then incubate the solution at 100 &#176;C for 10 min.</li>
	<li>Cool the heated solution to room temperature, add ultra-high purity urea powder into the solution to 8 M final concentration and incubate the sample at 37 &#176;C for 30 min.</li>
	<li>Add iodoacetamide stock solution to the sample to 15 mM final concentration, and incubate the solution in the dark for 30 min at room temperature to alkylate the free thiols in the sample. Add DTT stock solution to the sample to 10 mM final concentration and incubate the solution for another 10 min at room temperature to quench the excessive iodoacetamide.</li>
	<li>Dilute the obtained solution 10x&#160;with 40 mM Tris buffer at pH 8, and subsequently add trypsin to the sample at a 1:20 ratio of trypsin to total protein. Maintain the digestion reaction in a 37 &#176;C oven overnight to ensure the reaction is completed.</li>
	<li>Terminate the digestion by acidifying the sample solution to pH 1 with HCl, a condition that also breaks down the detergent, i.e. Rapigest. Degrade the Rapigest at 37 &#176;C for 1 hr, and remove the developed precipitation by centrifugation.</li>
	<li>Clean the supernatant that contains sample peptides by a C-18 solid phase extraction (SPE) cartridge and dry the obtained sample by speedvac.</li>
	<li>Perform a SDS-PAGE analysis of samples before and after trypsin digestion to confirm the digestion efficiency.</li>
</ol>
3. Glycopeptide Capture
<ol>
	<li>Dissolve the cleaned peptides in the coupling buffer (100 mM sodium acetate, pH 5.5). Add sodium periodate into the peptide solution at a 10 mM final concentration for 30 min in the dark, at room temperature. This will oxidize the cis-diol groups in the glycans to aldehydes, which allow the glycans to couple to the resin through hydrazide chemistry. Quench the excessive periodate by sodium sulphite at a 20 mM final concentration and pH 5 for 10 min at room temperature.</li>
	<li>Introduce hydrazide-derivatized resin into the peptide solution at a ratio of 1:4 (resin to solution) to couple glycopeptides to the resin. Incubate the reaction at 37 &#176;C for 1-2 days with end-to-end rotation for complete coupling.</li>
	<li>Remove the unbound peptides by washing the resin twice with 1 ml of each of the following: DI&#160;water, 1.5 M NaCl, methanol and 80% acetonitrile. Finally, wash the resin 3x&#160;with 1 ml of 100 mM NH4HCO3 at pH 8 to exchange the buffer of the system to 100 mM NH4HCO3.
	<ol>
		<li>Collect the supernatant and the washes for the analysis of unbound peptides (optional).</li>
	</ol>
	</li>
	<li>Release the N-glycopeptides from the resin by PNGase F in an overnight incubation at 37 &#176;C with an end-to-end rotation. Collect the released peptides by centrifugation and an 80% acetonitrile wash. Dry the obtained solution in the speedvac for LC-MS analysis.</li>
</ol>
4. Further Fractionation (Optional)
To further simplify sample complexity, fractionate the obtained N-glycopeptides. For example, redissolve the dried peptides into 10 mM ammonia formate, pH 3 with 20% acetonitrile and use strong&#160;cation&#160;exchange (SCX) chromatography to fractionate the peptides. Dry the obtained eluent, and then analyze the obtained peptide fractions by reverse-phase LC-MS8,17,18.
5. Cleaning of the Released N-glycopeptides (Optional)
If concerns rise for the potential contamination of the peptides, redissolve the dried peptides into 0.1% formic acid and use a MCX SPE column to further clean the peptides prior to reverse-phase LC-MS analysis.
Note: Database searching parameters.
During the selective cleavage of N-glycopeptides off the resin, PNGase F converts the N-glycan linked asparagine to an aspartic acid. Therefore, there is a 0.9840 Da mass shift of the liberated N-glycopeptides. To accurately identify these peptides, this modification needs to be added to the search parameters along with common modifications such as the carbamidomethylation of the cysteine and oxidation of the methionine.

<ol>
	<li>Cho, W., Stahelin, R. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane-protein+interactions+in+cell+signaling+and+membrane+trafficking.">Membrane-protein interactions in cell signaling and membrane trafficking.</a> Annual review of biophysics and biomolecular structure. 34, 119-151 (2005).</li><li>Tadevosyan, A., Vaniotis, G., Allen, B. G., Hebert, T. E., Nattel, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=protein-coupled+receptor+signalling+in+the+cardiac+nuclear+membrane:+evidence+and+possible+roles+in+physiological+and+pathophysiological+function.">protein-coupled receptor signalling in the cardiac nuclear membrane: evidence and possible roles in physiological and pathophysiological function.</a> The Journal of physiology. 590, 1313-1330 (2012).</li><li>White, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biophysical+dissection+of+membrane+proteins.">Biophysical dissection of membrane proteins.</a> Nature. 459, 344-346 (2009).</li><li>Heijne, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane-protein+topology.+.">Membrane-protein topology. .</a> Nature reviews Molecular cell biology. 7, 909-918 (2006).</li><li>Cox, J., Mann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+high-resolution+proteomics+for+data-driven+systems+biology.">Quantitative high-resolution proteomics for data-driven systems biology.</a> Annual review of biochemistry. 80, 273-299 (2011).</li><li>Lamond, A. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advancing+cell+biology+through+proteomics+in+space+and+time+(PROSPECTS).">Advancing cell biology through proteomics in space and time (PROSPECTS).</a> Mol Cell Proteomics. 11, (2012).</li><li>Savas, J. N., Stein, B. D., Wu, C. C., Yates 3rd, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mass+spectrometry+accelerates+membrane+protein+analysis.">Mass spectrometry accelerates membrane protein analysis.</a> Trends in biochemical sciences. 36, 388-396 (2011).</li><li>Sun, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shotgun+glycopeptide+capture+approach+coupled+with+mass+spectrometry+for+comprehensive+glycoproteomics.">Shotgun glycopeptide capture approach coupled with mass spectrometry for comprehensive glycoproteomics.</a> Mol Cell Proteomics. 6, 141-149 (2007).</li><li>Lowe, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycosylation,+immunity,+and+autoimmunity.">Glycosylation, immunity, and autoimmunity.</a> Cell. 104, 809-812 (2001).</li><li>Dodds, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gas-phase+dissociation+of+glycosylated+peptide+ions.">Gas-phase dissociation of glycosylated peptide ions.</a> Mass spectrometry reviews. 31, 666-682 (2012).</li><li>Kaji, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lectin+affinity+capture,+isotope-coded+tagging+and+mass+spectrometry+to+identify+N-linked+glycoproteins.">Lectin affinity capture, isotope-coded tagging and mass spectrometry to identify N-linked glycoproteins.</a> Nat Biotechnol. 21, 667-672 (2003).</li><li>Zhang, H., Li, X. J., Martin, D. B., Aebersold, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+quantification+of+N-linked+glycoproteins+using+hydrazide+chemistry,+stable+isotope+labeling+and+mass+spectrometry.">Identification and quantification of N-linked glycoproteins using hydrazide chemistry, stable isotope labeling and mass spectrometry.</a> Nat Biotechnol. 21, 660-666 (2003).</li><li>Hagglund, P., Bunkenborg, J., Elortza, F., Jensen, O. N., Roepstorff, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+strategy+for+identification+of+N-glycosylated+proteins+and+unambiguous+assignment+of+their+glycosylation+sites+using+HILIC+enrichment+and+partial+deglycosylation.">A new strategy for identification of N-glycosylated proteins and unambiguous assignment of their glycosylation sites using HILIC enrichment and partial deglycosylation.</a> J Proteome Res. 3, 556-566 (2004).</li><li>Bond, M. R., Kohler, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+methods+for+glycoprotein+discovery.">Chemical methods for glycoprotein discovery.</a> Current opinion in chemical biology. 11, 52-58 (2007).</li><li>Pasini, E. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In-depth+analysis+of+the+membrane+and+cytosolic+proteome+of+red+blood+cells.">In-depth analysis of the membrane and cytosolic proteome of red blood cells.</a> Blood. 108, 791-801 (2006).</li><li>Wollscheid, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mass-spectrometric+identification+and+relative+quantification+of+N-linked+cell+surface+glycoproteins.">Mass-spectrometric identification and relative quantification of N-linked cell surface glycoproteins.</a> Nat Biotechnol. 27, 378-386 (2009).</li><li>Sun, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=N-Glycoproteome+of+E14.Tg2a+mouse+embryonic+stem+cells.">N-Glycoproteome of E14.Tg2a mouse embryonic stem cells.</a> PLoS ONE. 8, (2013).</li><li>Sun, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycocapture-assisted+global+quantitative+proteomics+(gagQP)+reveals+multiorgan+responses+in+serum+toxicoproteome.">Glycocapture-assisted global quantitative proteomics (gagQP) reveals multiorgan responses in serum toxicoproteome.</a> J Proteome Res. 12, 2034-2044 (2013).</li></ol>

The authors declare that they have no competing financial interests. &#160;

Here we introduce a glycopeptide-capture strategy for profiling cell-surface proteins. The method can be applied to study secreted proteins, such as those in blood, as well as in other body fluids or in cell culture media.
The success of the method relies on the complete digestion of samples; therefore, a SDS-PAGE characterization of the digestion efficiency is necessary, especially for the first-time analysis of a sample. A complete digestion can be challenging for membrane proteins, and can only be possible after thorough solubilization of the membrane fraction. The solubilization process begins when the detergent is introduced and ends after the incubation of urea. Therefore, if cloudiness presents in the sample before the addition of urea but disappears after urea incubation, the solubilization is sufficient. If the membrane fraction is difficult to dissolve, increase the amount of detergent used. For membrane-rich tissue samples such as brain and adipose tissues, 5% Rapigest can be utilized. Sometimes, precipitation can appear during the dilution of samples prior to the trypsin digestion, which is more frequently observed for human serum samples. The cause of this precipitation is mainly the decreased concentrations of urea and detergent. This precipitation will generally be removed by trypsin during the digestion and is not a concern. However, when the precipitation forms, it is important to rotate the sample vial during the digestion step to ensure a good mixing. The pH of the glycocapture step is important because the primary amines in peptides, such as those from N-termini and lysines, will react with the newly formed aldehydes after the periodate oxidation. Thus, it is important to protonate these amines by adjusting the pH of the capturing solution to below 6.0 using acetate buffer, to prevent them from interfering with the capture.
Using a ratio of resin to capture solution around 1:3 to 1:4 ensures sufficient mixing during the capture step. A minimum of 50 &#956;l of resin is necessary based on our experience; lower quantities will render the subsequent series of washes difficult to perform and will introduce severe sample loss due to the loss of resin. We have obtained good results using 100-300 &#956;l of resin for 0.5-2 mg of total protein. However, the ratio between the amount of protein and resin can be sample dependent, we recommend you optimize this condition for your specific samples and applications.
The hydrazide chemistry captures both O- and N-linked glycopeptides on the substrate; due to the lack of an effective enzyme to release all the O-linked glycopeptides, we only used PNGase F for N-glycopeptide studies8,12. The possibility of studying the O-type of glycopeptides may require the discovery or bioengineering of appropriate hydrolases.
This method can be paused at several places including: 1) after obtaining the membrane fraction, 2) after trypsin digestion and cleaning of the peptides, and 3) after release of the N-glycopeptides. Additional procedures can be introduced during these intervals. For example, the digested peptides can be differentially labeled by N-isotags as indicated in Figure 1, for quantitative analysis. Using a method called gagQP, in which the unmodified peptides are analyzed in parallel with the glycopeptides, the accuracy of quantitation can be significantly improved as we demonstrated in a recent publication18.
Glycopeptide capture itself can effectively decrease sample complexity, and it is generally not necessary to further fractionate the enriched N-glycopeptides. Exceptions apply to situations where the samples are abundant but with substantial concentration dynamics, and the proteins of interest are in low abundance, such as in the discovery of blood protein biomarkers or for a complete survey of cell surface proteins. Under those circumstances further fractionation can be implemented for the captured N-glycopeptides to provide a better penetration of the glycoproteome. As the bottom of the glycoproteome is being approached through fractionation, the identification of non-glycopeptides introduced by nonspecific binding will increase. Thus a decrease of glycopeptide and glycoprotein selectivity (to ~85%) will typically be observed after a further fractionation of N-glycopeptides17. Therefore, researchers need to weigh the pros and cons when designing the most suitable procedure.
For researchers who have never used an N-glycopeptide-capture method, it is best to practice the method with a few pure N-glycoprotein(s) having known glycosylation sites as listed previously8. This method is robust and the selectivity to N-glycopeptides is high. A drawback of this method lies in the inherent low abundance of surface N-glycoproteins; for a comprehensive characterization of the sample, higher quantity is generally required than that of cytosolic proteomics. However, if cultured cells can be expanded in vitro or the tissues of interest are abundant, this drawback is negligible.

Cell surface proteins interact with the extracellular environment and relay signals from the outside to the inside of a cell. Thus, these proteins,&#160;including extracellular matrix proteins,&#160;play critical roles in all aspects of cellular biology and physiology ranging from proliferation, growth, migration, differentiation to aging and so forth. Surface proteins function by interacting with other cells, proteins and small molecules1-3. Molecular characterization of cell-surface proteins is of great interest not only for biologists but also for pharmaceutical companies, as more than 60% of drugs are targeted to cell-surface proteins4.
Tandem mass spectrometry (MS), with its superior sensitivity, accuracy, and throughput for identification of proteins and peptides, has been a powerful tool for global proteomic studies5,6. Yet, surface proteins pose significant challenges to MS-based proteomics, as most surface proteins exist in low quantities and with heavy modifications. The membrane-spanning regions of the surface proteins render them hydrophobic; this is especially the case for multipass transmembrane proteins. It is thus difficult to dissolve membrane proteins in aqueous solutions without the help of a detergent; however the use of detergents generally suppresses the performance of HPLC and MS1,7,8 in protein identification. Therefore, membrane proteins have been poorly characterized in direct LC-MS based proteomics.
Glycosylation is one of the most important and abundant post-translational modifications taking place in cell-surface proteins9. The enormous complexity and heterogeneity of glycans hamper peptides&#8217; MS signal10. Nevertheless, several proteomic methods have used this unique modification to enrich surface proteins and to remove the sugar moieties from proteins prior to LC-MS analysis. These methods include lectin-based affinity capture11 and hydrazide-based or boric acid-based chemical capture12 as well as hydrophilic chromatography separations8,13. The removal of glycans transforms membrane proteins to regular proteins and drastically simplifies the MS characterization. Because glycosylation also takes place in secreted proteins that have high solubility in contrast to membrane proteins, many glycoproteomic methods are optimized for soluble proteins, and tend to have lower glycopeptide selectivity and sensitivity when being deployed to membrane proteins8,14. Other methods also exist to enrich, in particular, cell-surface proteins, such as those using ultracentrifugation15 and labeling strategies16. A detailed comparison between our method and other existing methods for characterizing membrane proteins was conducted recently17, and the results indicated that our method can perform equally well, if not better, than all the compared membrane proteomics methods, but with higher simplicity.
To help researchers use this method, we detail here a general protocol. This method integrates several advantages of existing glycoproteomics strategies and is devised specifically for membrane glycoproteins, yet the method works equally well for secreted proteins. The characteristics of this method include: 1) a complete solubilization of membrane proteins, 2) an enrichment of glycopeptides instead of glycoproteins to eliminate the potential steric hindrance when using a solid capturing substrate, 3) the use of hydrazide chemistry to form covalent bonds between glycopeptides and the capturing substrate, such that the bonded glycopeptides can tolerate stringent washes for high glycoselectivity, and 4) the capability to conduct the entire capture procedure in one tube for reduced sample loss and shortened procedure duration. After implementing this method to studying a variety of biological samples including cells and tissues, we observed a high selectivity (&#62; 90%) to glycoproteins8,17,18.

<table border="0" cellpadding="0" cellspacing="0" width="701">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DTT</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">646563</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">TCEP</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">646547</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Iodoacetamide</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">A3221</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rapigest SF</td>
			<td style="width:199px;">Waters</td>
			<td style="width:119px;">186001860</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium periodate</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">311448</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PNGase F</td>
			<td style="width:199px;">New England Biolabs</td>
			<td style="width:119px;">P0704S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Affi-Gel Hz Hydrazide gel</td>
			<td style="width:199px;">Bio-Rad</td>
			<td style="width:119px;">153-6047</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Trypsin</td>
			<td style="width:199px;">Worthington Biomedical</td>
			<td style="width:119px;">LS02115</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sep-Pak C-18 cartridge</td>
			<td style="width:199px;">Waters</td>
			<td style="width:119px;">WAT054955</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Oasis MCX cartridge</td>
			<td style="width:199px;">Waters</td>
			<td style="width:119px;">186000252</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Protease inhibitor coctail</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">P8340</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Urea</td>
			<td style="width:199px;">Amersco</td>
			<td style="width:119px;">568</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium sulphite</td>
			<td style="width:199px;">Caledon</td>
			<td style="width:119px;">8360-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Invertase</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">I0408</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Alpha-1 trypsin</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">F2006</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ribonulease B</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">R7884</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Avidin</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">A9275</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ovalbumin</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">A5503</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Conalbumin</td>
			<td style="width:199px;">Sigma</td>
			<td style="width:119px;">C0755</td>
			<td></td>
		</tr>
	</tbody>
</table>

glycopeptide capture for cell surface proteomics

Cell surface proteins, including extracellular matrix proteins, participate in all major cellular processes and functions, such as growth, differentiation, and proliferation. A comprehensive characterization of these proteins provides rich information for biomarker discovery, cell-type identification, and drug-target selection, as well as helping to advance our understanding of cellular biology and physiology. Surface proteins, however, pose significant analytical challenges, because of their inherently low abundance, high hydrophobicity, and heavy post-translational modifications. Taking advantage of the prevalent glycosylation on surface proteins, we introduce here a high-throughput glycopeptide-capture approach that integrates the advantages of several existing N-glycoproteomics means. Our method can enrich the glycopeptides derived from surface proteins and remove their glycans for facile proteomics using LC-MS. The resolved N-glycoproteome comprises the information of protein identity and quantity as well as their sites of glycosylation. This method has been applied to a series of studies in areas including cancer, stem cells, and drug toxicity. The limitation of the method lies in the low abundance of surface membrane proteins, such that a relatively large quantity of samples is required for this analysis compared to studies centered on cytosolic proteins.

A representative flow chart of the experimental procedure is summarized in Figure 1. The labeling and further fractionation steps are optional and details are described in a recent publication18. Another option is to analyze the unmodified peptides, which do not react with the resin. The advantages of analyzing the unmodified peptides include the potential identification of non-glycosylated peptides and proteins, such as claudins in tight junctions; an additional advantage is more accurate quantitation. Based on these advantages, we termed this method glycocapture-assisted-global quantitative proteomics (gagQP), and detailed the analysis in a recent publications18.
A typical glycopeptide spectrum taken after the enrichment method is shown in Figure 2. In the obtained glycopeptide, the N-glycan was removed and the glycan-attached asparagine (N) was converted to an aspartic acid (D) by PNGase F; therefore, the spectrum can be readily searched using any proteomics search engine against common protein sequence databases.
The capture method can be evaluated by commercially available model glycoproteins prior to its application with complex and valuable biological samples. Some frequently used model glycoproteins include avidin (chicken), ovalbumin (chicken), invertase (yeast), &#945;-1 antitrypsin (human), conalbumin (chicken), and ribonuclease B (bovine) (all can be obtained from Sigma). The frequently identified glycopeptides from these proteins can be found in a previous publication8. A customized protein-sequence FASTA&#160;database that is suitable for automatic searching of LC-MS results generated from these model proteins can be downloaded from the following link (http://www.sfu.ca/chemistry/groups/bingyun_sun/tools.html), which includes the above listed model proteins as well as a reversed yeast-sequence database, common contaminants and PNGase F.
A typical LC-MS result of the captured N-glycopeptides is shown in Figure 3, in which more than 100 glycoproteins can be identified from a single LC-MS run of a cell microsomal fraction. The enrichment selectivity to both glycoproteins and glycopeptides is generally more than 90%. A successful analysis can have an enrichment selectivity of 95%. Using an SCX column and step gradient to further fractionate the samples prior to LC-MS analyses will usually double the number of glycoprotein identifications17. Sometimes, when the quantity of the obtained glycopeptides is low, the impurity accumulated from the vials can be observed in the final sample. These contaminants can be removed by the method provided in step 5.
<img alt="Figure 1" fo:content-width="3in" fo:src="/files/ftp_upload/51349/51349fig1highres.jpg" src="/files/ftp_upload/51349/51349fig1.jpg" width="350" /> 
Figure 1.&#160;Flow chart of the experimental procedure. Rectangles are required steps and diamonds are optional steps.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51349/51349fig2highres.jpg" src="/files/ftp_upload/51349/51349fig2.jpg" /> 
Figure 2.&#160;Collision induced dissociation (CID) spectrum of AEPILNISNAGPWSR from Baker&#8217;s yeast after the glycopeptide-capture method. The underscored asparagine (N) is converted to the aspartic acid (D) as highlighted in red in the peptide sequence in the figure.
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51349/51349fig1h3ghres.jpg" src="/files/ftp_upload/51349/51349fig3.jpg" /> 
Figure 3.&#160;2D plot of the LC-MS result of the N-glycopeptides obtained from mouse embryonic stem cells (E14.Tg2a). The dots are the detected peptides, and the color of the dot represents the identification confidence obtained statistically (i.e. peptide probability).

Watch this Scientific Journal Video about Glycopeptide Capture for Cell Surface Proteomics at JoVE.com

Glycopeptide Capture for Cell Surface Proteomics

Cell surface proteins are biologically important and widely glycosylated. We introduce here a glycopeptide-capture approach to solubilize, enrich, and deglycosylate these proteins for facile LC-MS based proteomic analyses.

Cell surface proteins, including extracellular matrix proteins, participate in all major cellular processes and functions, such as growth, ...

glycopeptide-capture-for-cell-surface-proteomics

Nanyang Technological University

Research

JoVE Journal

Biology

11.2K Views.  Simon Fraser University. Cell surface proteins are biologically important and widely glycosylated. We introduce here a glycopeptide-capture approach to solubilize, enrich, and deglycosylate these proteins for facile LC-MS based proteomic analyses.

Video: Glycopeptide Capture for Cell Surface Proteomics

Cell Capture Using a Microfluidic Device

Orthogonal Protein Purification Facilitated by a Small Bispecific Affinity Tag

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a demand for general methods that do not require target protein specific optimization1 . To achieve this, purification tags that genetically can be fused to the gene of interest are commonly used2 . The most widely used affinity handle is the hexa-histidine tag, which is suitable for purification under both native and denaturing conditions3 . The metabolic burden for producing the tag is low, but it does not provide as high specificity as competing affinity chromatography based strategies1,2.
Here, a bispecific purification tag with two different binding sites on a 46 amino acid, small protein domain has been developed. The albumin-binding domain is derived from Streptococcal protein G and has a strong inherent affinity to human serum albumin (HSA). Eleven surface-exposed amino acids, not involved in albumin-binding4 , were genetically randomized to produce a combinatorial library. The protein library with the novel randomly arranged binding surface (Figure 1) was expressed on phage particles to facilitate selection of binders by phage display technology. Through several rounds of biopanning against a dimeric Z-domain derived from Staphylococcal protein A5, a small, bispecific molecule with affinity for both HSA and the novel target was identified6 .
The novel protein domain, referred to as ABDz1, was evaluated as a purification tag for a selection of target proteins with different molecular weight, solubility and isoelectric point. Three target proteins were expressed in Escherishia coli with the novel tag fused to their N-termini and thereafter affinity purified. Initial purification on either a column with immobilized HSA or Z-domain resulted in relatively pure products. Two-step affinity purification with the bispecific tag resulted in substantial improvement of protein purity. Chromatographic media with the Z-domain immobilized, for example MabSelect SuRe, are readily available for purification of antibodies and HSA can easily be chemically coupled to media to provide the second matrix.
This method is especially advantageous when there is a high demand on purity of the recovered target protein. The bifunctionality of the tag allows two different chromatographic steps to be used while the metabolic burden on the expression host is limited due to the small size of the tag. It provides a competitive alternative to so called combinatorial tagging where multiple tags are used in combination1,7.

A novel and highly efficient two-step affinity chromatography protocol has been developed and is described in detail. The method is based on a small purification tag with two inherent affinities and is applicable to a wide range of target proteins with different properties.

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Avidity-based Extracellular Interaction Screening (AVEXIS) for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ensuring cohesion within multicellular organisms. Proteins predicted to form extracellular interactions are encoded by approximately a quarter of human genes1, but despite their importance and abundance, the majority of these proteins have no documented binding partner. Primarily, this is due to their biochemical intractability: membrane-embedded proteins are difficult to solubilise in their native conformation and contain structurally-important posttranslational modifications. Also, the interaction affinities between receptor proteins are often characterised by extremely low interaction strengths (half-lives &lt; 1 second) precluding their detection with many commonly-used high throughput methods2. 
Here, we describe an assay, AVEXIS (AVidity-based EXtracellular Interaction Screen) that overcomes these technical challenges enabling the detection of very weak protein interactions (t1/2 &le; 0.1 sec) with a low false positive rate3. The assay is usually implemented in a high throughput format to enable the systematic screening of many thousands of interactions in a convenient microtitre plate format (Fig. 1). It relies on the production of soluble recombinant protein libraries that contain the ectodomain fragments of cell surface receptors or secreted proteins within which to screen for interactions; therefore, this approach is suitable for type I, type II, GPI-linked cell surface receptors and secreted proteins but not for multipass membrane proteins such as ion channels or transporters.
The recombinant protein libraries are produced using a convenient and high-level mammalian expression system4, to ensure that important posttranslational modifications such as glycosylation and disulphide bonds are added. Expressed recombinant proteins are secreted into the medium and produced in two forms: a biotinylated bait which can be captured on a streptavidin-coated solid phase suitable for screening, and a pentamerised enzyme-tagged (&beta;-lactamase) prey. The bait and prey proteins are presented to each other in a binary fashion to detect direct interactions between them, similar to a conventional ELISA (Fig. 1). The pentamerisation of the proteins in the prey is achieved through a peptide sequence from the cartilage oligomeric matrix protein (COMP) and increases the local concentration of the ectodomains thereby providing significant avidity gains to enable even very transient interactions to be detected. By normalising the activities of both the bait and prey to predetermined levels prior to screening, we have shown that interactions having monomeric half-lives of 0.1 sec can be detected with low false positive rates3.

Avidity-based Extracellular Interaction Screening AVEXIS for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

AVEXIS is a high throughput protein interaction assay developed to systematically screen for novel extracellular receptor-ligand pairs involved in cellular recognition processes. It is specifically designed to detect transient protein interactions that are difficult to identify using other high throughput approaches.

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ...

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 ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis (diguanylate cyclases) and degradation (phosphodiesterases) of the second messenger c-di-GMP. In contrast, little information is available regarding the molecular mechanisms and cellular components through which this signaling molecule regulates a diverse range of cellular processes. Most of the known effector proteins belong to the PilZ family or are degenerated diguanylate cyclases or phosphodiesterases that have given up on catalysis and have adopted effector function. Thus, to better define the cellular c-di-GMP network in a wide range of bacteria experimental methods are required to identify and validate novel effectors for which reliable in silico predictions fail.
We have recently developed a novel Capture Compound Mass Spectrometry (CCMS) based technology as a powerful tool to biochemically identify and characterize c-di-GMP binding proteins. This technique has previously been reported to be applicable to a wide range of organisms1. Here we give a detailed description of the protocol that we utilize to probe such signaling components. As an example, we use Pseudomonas aeruginosa, an opportunistic pathogen in which c-di-GMP plays a critical role in virulence and biofilm control. CCMS identified 74% (38/51) of the known or predicted components of the c-di-GMP network. This study explains the CCMS procedure in detail, and establishes it as a powerful and versatile tool to identify novel components involved in small molecule signaling.

The ubiquitous second messenger c-di-GMP controls growth and behavior of many bacteria. We have developed a novel Capture Compound Mass Spectrometry based technology to biochemically identify and characterize c-di-GMP binding proteins in virtually any bacterial species.

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis ...

Cell Surface Marker Mediated Purification of iPS Cell Intermediates from a Reprogrammable Mouse Model

Mature cells can be reprogrammed to a pluripotent state. These so called induced pluripotent stem (iPS) cells are able to give rise to all cell types of the body and consequently have vast potential for regenerative medicine applications. Traditionally iPS cells are generated by viral introduction of transcription factors Oct-4, Klf-4, Sox-2, and c-Myc (OKSM) into fibroblasts. However, reprogramming is an inefficient process with only 0.1-1% of cells reverting towards a pluripotent state, making it difficult to study the reprogramming mechanism. A proven methodology that has allowed the study of the reprogramming process is to separate the rare intermediates of the reaction from the refractory bulk population. In the case of mouse embryonic fibroblasts (MEFs), we and others have previously shown that reprogramming cells undergo a distinct series of changes in the expression profile of cell surface markers which can be used for the separation of these cells. During the early stages of OKSM expression successfully reprogramming cells lose fibroblast identity marker Thy-1.2 and up-regulate pluripotency associated marker Ssea-1. The final transition of a subset of Ssea-1 positive cells towards the pluripotent state is marked by the expression of Epcam during the late stages of reprogramming. Here we provide a detailed description of the methodology used to isolate reprogramming intermediates from cultures of reprogramming MEFs. In order to increase experimental reproducibility we use a reprogrammable mouse strain that has been engineered to express a transcriptional transactivator (m2rtTA) under control of the Rosa26 locus and OKSM under control of a doxycycline responsive promoter. Cells isolated from these mice are isogenic and express OKSM homogenously upon addition of doxycycline. We describe in detail the establishment of the reprogrammable mice, the derivation of MEFs, and the subsequent isolation of intermediates during reprogramming into iPS cells via fluorescent activated cells sorting (FACS).

Mouse embryonic fibroblast can be reprogrammed into induced pluripotent stem cells at low efficiency by the forced expression of transcription factors Oct-4, Sox-2, Klf-4, c-Myc. The rare intermediates of the reprogramming reaction are FACS isolated via labeling with antibodies against cell surface makers Thy-1.2, Ssea-1, and Epcam.

Mature cells can be reprogrammed to a pluripotent state. These so called induced pluripotent stem (iPS) cells are able to give rise to all cell types of ...

Non-enzymatic, Serum-free Tissue Culture of Pre-invasive Breast Lesions for Spontaneous Generation of Mammospheres

Breast ductal carcinoma in situ (DCIS), by definition, is proliferation of neoplastic epithelial cells within the confines of the breast duct, without breaching the collagenous basement membrane. While DCIS is a non-obligate precursor to invasive breast cancers, the molecular mechanisms and cell populations that permit progression to invasive cancer are not fully known. To determine if progenitor cells capable of invasion existed within the DCIS cell population, we developed a methodology for collecting and culturing sterile human breast tissue at the time of surgery, without enzymatic disruption of tissue.
Sterile breast tissue containing ductal segments is harvested from surgically excised breast tissue following routine pathological examination. Tissue containing DCIS is placed in nutrient rich, antibiotic-containing, serum free medium, and transported to the tissue culture laboratory. The breast tissue is further dissected to isolate the calcified areas. Multiple breast tissue pieces (organoids) are placed in a minimal volume of serum free medium in a flask with a removable lid and cultured in a humidified CO2 incubator. Epithelial and fibroblast cell populations emerge from the organoid after 10 - 14 days. Mammospheres spontaneously form on and around the epithelial cell monolayer. Specific cell populations can be harvested directly from the flask without disrupting neighboring cells. Our non-enzymatic tissue culture system reliably reveals cytogenetically abnormal, invasive progenitor cells from fresh human DCIS lesions.

Primary cell culture using intact tissue organoids provides a model system that mimics the multi-cellular in vivo microenvironment. We developed a serum-free primary breast epithelium tissue culture model that perpetuates mixed cell culture lineages and exhibits differentiated morphology, without enzymatic tissue disruption. Breast organoids remain viable for &#62;6 months.

Breast ductal carcinoma in situ (DCIS), by definition, is proliferation of neoplastic epithelial cells within the confines of the breast duct, without ...

Selected Reaction Monitoring Mass Spectrometry for Absolute Protein Quantification

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This protocol provides step-by-step instructions for the development and application of quantitative assays using selected reaction monitoring (SRM) mass spectrometry (MS). First, likely quantotypic target peptides are identified based on numerous criteria. This includes identifying proteotypic peptides, avoiding sites of posttranslational modification, and analyzing the uniqueness of the target peptide to the target protein. Next, crude external peptide standards are synthesized and used to develop SRM assays, and the resulting assays are used to perform qualitative analyses of the biological samples. Finally, purified, quantified, heavy isotope labeled internal peptide standards are prepared and used to perform isotope dilution series SRM assays. Analysis of all of the resulting MS data is presented. This protocol was used to accurately assay the absolute abundance of proteins of the chemotaxis signaling pathway within RAW 264.7 cells (a mouse monocyte/macrophage cell line). The quantification of Gi2 (a heterotrimeric G-protein &#945;-subunit) is described in detail.

This protocol describes how to perform absolute quantification assays of target proteins within complex biological samples using selected reaction monitoring. It was used to accurately quantify proteins of the mouse macrophage chemotaxis signaling pathway. Target peptide selection, assay development, and qualitative and quantitative assays are described in detail.

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...