This work was funded by the NCI Clinical Proteomic Technology Assessment Center (CPTAC) grant (#U24 CA126476) as well as a grant from the Entertainment Industry Foundation (EIF) and the EIF Women's Cancer Research Fund to the Breast Cancer Biomarker Discovery Consortium, and generous gifts from the Keck Foundation, the Canary Foundation, and the Paul G. Allen Family Foundation.

Experimental Procedure : 
The assay requires synthetic peptides and anti-peptide antibodies. Selected peptides should be unique to the protein of interest, contain between 8 and 22 amino acids, and have no known post-translational modifications. Methionine residues are generally avoided and peptides containing dibasic amino acids (e.g. KK, KR, RR) are undesirable. For this technique, it is common to use stable isotope labeled peptides as internal standards, incorporating heavy (13C and 15N) labeled amino acids at the C-terminus of the peptide (i.e. K or R labeled). 
The following protocol describes an assay developed to measure the peptide GDSLAYGLR from the mouse protein Osteopontin, using anti-peptide antibodies obtained from Epitomics Inc. (Burlingame, CA) and synthetic peptides from New England Peptide (Gardner, MA). The protocol consists of three main steps (Figure 1): 1) Trypsin digestion of the complex protein mixture, 2) Enrichment of peptides 3) Analysis by mass spectrometry. It will be demonstrated on a human plasma sample spiked with the mouse Osteopontin protein. 
1. Trypsin enzymatic digestion and cleanup
<ol>
<li>Thaw 10 &mu;L neat plasma aliquot on wet ice.</li>
<li>Determine the total protein concentration by BCA assay and centrifuge the sample to remove any suspended solids.</li>
<li>Pipet 10 &mu;L aliquot from its storage tube to a 1000 &mu;L deepwell plate and cover with pierce-able film.</li>
<li>Add 20 &mu;L of fresh 9M urea / 30mM dithiothreitol (DTT) (final concentration 6M urea / 20mM DTT) to each sample.</li>
<li>Incubate for 30 minutes at 37&deg;C.</li>
<li>Add 3 &mu;L of fresh 500 mM iodoacetamide (final IAM 50mM) to each sample.</li>
<li> Incubate for 30 minutes in the dark at room temperature.</li>
<li>Add 257 &mu;L of 100 mM Tris (pH 8) (dilutes urea to &tilde;0.6M).</li>
<li>Add 10 &mu;L of trypsin stock solution (1 &mu;g/&mu;L; for 1:50 enzyme:substrate ratio).</li>
<li>Incubate 37&deg;C overnight (12-16 hours).</li>
<li>Add 3 &mu;L of neat formic acid (final concentration of 1%).</li>
<li>Add stable isotope standard (multiple standards are added if performing a multiplexed assay, typically this is about 10 &mu;L containing 50-100 fmol of standard isotopically-labeled peptide).</li>
<li> Wash the Oasis cartridge plate well with 500 &mu;L of 0.1% formic acid in 80% acetonitrile, discarding the flow-through. Repeat this 3 times.</li>
<li>Equilibrate the cartridge plate by adding 500 &mu;L of 0.1% formic acid in water, and discard the flow-through. Repeat this 4 times.</li>
<li>Load digest samples to the cartridge plate and adjust the vacuum so the flow is very slow.</li>
<li>Wash with 500 &mu;L of 0.1% formic acid in water, and discard flow-through. Repeat this 3 times.</li>
<li>Elute peptides by adding 2 x 500 &mu;L of 0.1% formic acid in 80% acetonitrile into 1000 &mu;l deep-well plate (do not discard the flow-through).</li>
<li>Lyophilize (or speedvac) the eluate to dryness. (Lyophilization is the preferred method)</li>
<li>Reconstitute dried peptides by adding 50 &mu;L PBS + 0.03% CHAPS.</li> 
</ol>
2. Peptide immunoaffinity enrichment
<ol>
<li>Transfer the sample to standard Kingfisher 96 well plates.</li>
<li>Add 1 &mu;g antibody and 1.5 &mu;L Protein-G coated magnetic beads per target (it is optional to crosslink the antibody to the beads prior to addition). Ensure beads are well suspended by shaking or vortexing.</li>
<li>Cover the plate with foil.</li>
<li>Incubate overnight (12-16 hours) with gentle tumbling to ensure beads are suspended.</li>
<li>Centrifuge plate at 32 x g for 5 seconds to remove any liquid from the foil surface.</li>
<li>Remove the foil cover.</li>
<li>Washing and elution of the beads can be performed manually or in an automated fashion. This procedure describes the steps automated on a Kingfisher platform.</li>
<li>Wash the beads 2 times with 200 &mu;L PBS + 0.03% CHAPS (1 minute per wash).</li>
<li>Wash the beads 1 time with 200 &mu;L 1:10 dilution of PBS + 0.03% CHAPS (1 minute).</li>
<li>The peptides are eluted in 25 uL of 5% acetic acid + 0.03% CHAPS.</li>
<li>Place the elution plate on a magnet and transfer the eluate to a 96 well plate, taking care not to transfer the leftover beads.</li>
<li>Cover the plate with a sealing mat.</li>
<li>The plate containing the eluate is delivered to the triple quadrapole mass spectrometer for analysis.</li>
</ol>
3. Analysis by multiple reaction monitoring - mass spectrometry
<ol>
<li>Transitions for SRM/MRM analysis can be selected and optimized by infusing a 1 picomole per microliter solution of peptide standard in 30% acetonitrile/0.1% formic acid into the mass spectrometer at 0.5 &mu;L/min. Once stable spray is obtained, collect an MS/MS spectrum. </li>
<li>Transitions are selected from the MS/MS spectrum by identifying three abundant fragment ions in a region of the spectrum with little noise. For multiply-charged peptide ions, the y-ion fragments detected at m/z &gt; precursor m/z are typically the best for SRM/MRM assay development. Figure 2 shows the MS/MS spectrum and selected transition ions 476.3 &gt; 508.3, 579.3, 692.4 (transitions for the heavy stable isotope standard 481.3 &gt; 518.3, 589.3, 702.4 not shown) for the peptide GDSLAYGLR. </li>
<li>Depending on the make and model of the mass spectrometer used, there are multiple parameters that can be optimized for each transition. Here, we optimized the collision energy of each selected transition by ramping the collision energy and monitoring the level of signal. A collision energy of 25 was used for each transition. </li>
<li>Briefly, a typical configuration for SRM/MRM analysis is as follows: Mobile phase (A) 0.1% Formic acid; (B) 90% Acetonitrile / 0.1% Formic acid, 0.3 x 5 mm C18 trap column, 75 &mu;m ID x 15 cm C18 analytical column (Reprosil-Pur C18 AQ, 120 A&deg; pore). The injection volume is 10 &mu;L and the sample is loaded for 5 min at 3% B at a total flow rate 3 &mu;L/min and eluted by a linear gradient from 3 - 45 %B at 300 - 400 nL/min in 10 min. Conditions on a 4000 QTRAP (ABSCIEX, Foster City, CA) were spray voltage 2.3kV, ion source temperature 150 &deg;C, GS1 of 12, and curtain gas 15. </li>
<li>The sample is analyzed by SRM/MRM by injecting 10 &mu;L of the sample. Peak areas are integrated for light and heavy peptides using the program Skyline.14 Calculate the peak area ratio (PAR) of unlabeled/labeled peptide in each sample using the most abundant transition that is free of background noise, in this case, the y5 transition (light peptide 476.3 &gt; 579.3, heavy standard peptide 481.3 &gt; 589.3). </li>
</ol>
4. Representative Results: 
Measured peak area ratios (light endogenous peptide relative to spiked heavy isotopically-labeled peptide) provide a quantitative measure of the target peptide. Figure 3 shows an example chromatogram of light and heavy peptides in a SISCAPA-enriched sample. Note that light and heavy peptides elute at the same time and multiple transitions can be monitored for each peptide to confirm the identity.
<img src="/files/ftp_upload/2812/2812fig1.jpg" alt="Figure 1"/> 
Figure 1. A schematic overview of the SISCAPA process. A complex protein mixture is digested into peptides. Targeted peptide analytes (endogenous analyte and a spiked stable isotope-labeled internal standard) are enriched using anti-peptide antibodies immobilized on Protein G-coated magnetic particles. Following isolation, the targeted peptides are eluted from the magnetic particles and analyzed by mass spectrometry for quantitation relative to the internal standard.
<img src="/files/ftp_upload/2812/2812fig2.jpg" alt="Figure 2"/> 
Figure 2. MS/MS spectrum of the peptide GDSLAYGLR showing selection of three fragments for SRM/MRM transitions.

<img src="/files/ftp_upload/2812/2812fig3.jpg" alt="Figure 3"/> 
Figure 3. Example chromatograms showing the peak profile of a light peptide analyte (red) and the heavy stable isotope-labeled internal standard (blue). Chromatograms for the y5 transition from the light peptide (476.3 &gt; 579.3) and heavy stable isotope labeled standard (481.3 &gt; 589.3) are plotted over time as they elute from the chromatography system.

<ol>
	<li>Hoofnagle, A. N., Wener, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+fundamental+flaws+of+immunoassays+and+potential+solutions+using+tandem+mass+spectrometry.">The fundamental flaws of immunoassays and potential solutions using tandem mass spectrometry.</a> J. Immunol. Methods. 347, 3-11 (2009).</li><li>Anderson, N. L. <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+quantitation+of+peptides+and+proteins+using+Stable+Isotope+Standards+and+Capture+by+Anti-Peptide+Antibodies+(SISCAPA).">Mass spectrometric quantitation of peptides and proteins using Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA).</a> J Proteome Res. 3, 235-244 (2004).</li><li>Chace, D. H., Kalas, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+biochemical+perspective+on+the+use+of+tandem+mass+spectrometry+for+newborn+screening+and+clinical+testing.">A biochemical perspective on the use of tandem mass spectrometry for newborn screening and clinical testing.</a> Clin Biochem. 38, 296-309 (2005).</li><li>Want, E. J., Cravatt, B. F., Siuzdak, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+expanding+role+of+mass+spectrometry+in+metabolite+profiling+and+characterization.">The expanding role of mass spectrometry in metabolite profiling and characterization.</a> Chembiochem. 6, 1941-1951 (2005).</li><li>Barr, 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=Isotope+dilution--mass+spectrometric+quantification+of+specific+proteins:+model+application+with+apolipoprotein+A-I.">Isotope dilution--mass spectrometric quantification of specific proteins: model application with apolipoprotein A-I.</a> Clin Chem. 42, 1676-1682 (1996).</li><li>Gerber, S. A., Rush, J., Stemman, O., Kirschner, M. W., Gygi, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Absolute+quantification+of+proteins+and+phosphoproteins+from+cell+lysates+by+tandem+MS.">Absolute quantification of proteins and phosphoproteins from cell lysates by tandem MS.</a> Proc Natl Acad Sci U S A. 100, 6940-6945 (2003).</li><li>Kuhn, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+C-reactive+protein+in+the+serum+of+patients+with+rheumatoid+arthritis+using+multiple+reaction+monitoring+mass+spectrometry+and+13C-labeled+peptide+standards.">Quantification of C-reactive protein in the serum of patients with rheumatoid arthritis using multiple reaction monitoring mass spectrometry and 13C-labeled peptide standards.</a> Proteomics. 4, 1175-1186 (2004).</li><li>Whiteaker, 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=Antibody-based+enrichment+of+peptides+on+magnetic+beads+for+mass-spectrometry-based+quantification+of+serum+biomarkers.">Antibody-based enrichment of peptides on magnetic beads for mass-spectrometry-based quantification of serum biomarkers.</a> Anal Biochem. 362, 44-54 (2007).</li><li>Hoofnagle, A. N., Becker, J. O., Wener, M. H., Heinecke, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+thyroglobulin,+a+low-abundance+serum+protein,+by+immunoaffinity+peptide+enrichment+and+tandem+mass+spectrometry.">Quantification of thyroglobulin, a low-abundance serum protein, by immunoaffinity peptide enrichment and tandem mass spectrometry.</a> Clin Chem. 54, 1796-1804 (2008).</li><li>Kuhn, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developing+Multiplexed+Assays+for+Troponin+I+and+Interleukin-33+in+Plasma+by+Peptide+Immunoaffinity+Enrichment+and+Targeted+Mass+Spectrometry.">Developing Multiplexed Assays for Troponin I and Interleukin-33 in Plasma by Peptide Immunoaffinity Enrichment and Targeted Mass Spectrometry.</a> Clin Chem. 55, 1108-1117 (2009).</li><li>Whiteaker, J. R., Zhao, L., Anderson, L., Paulovich, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+automated+and+multiplexed+method+for+high+throughput+peptide+immunoaffinity+enrichment+and+multiple+reaction+monitoring+mass+spectrometry-based+quantification+of+protein+biomarkers.">An automated and multiplexed method for high throughput peptide immunoaffinity enrichment and multiple reaction monitoring mass spectrometry-based quantification of protein biomarkers.</a> Mol. Cell. Proteomics. 9, 184-196 (2010).</li><li>Neubert, H., Gale, J., Muirhead, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Online+high-flow+peptide+immunoaffinity+enrichment+and+nanoflow+LC-MS/MS:+assay+development+for+total+salivary+pepsin/pepsinogen.">Online high-flow peptide immunoaffinity enrichment and nanoflow LC-MS/MS: assay development for total salivary pepsin/pepsinogen.</a> Clin. Chem. 56, 1413-1423 (2010).</li><li>Ackermann, B. L., Berna, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coupling+immunoaffinity+techniques+with+MS+for+quantitative+analysis+of+low-abundance+protein+biomarkers.">Coupling immunoaffinity techniques with MS for quantitative analysis of low-abundance protein biomarkers.</a> Expert Rev. Proteomics. 4, 175-186 (2007).</li><li>MacLean, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skyline:+an+open+source+document+editor+for+creating+and+analyzing+targeted+proteomics+experiments.">Skyline: an open source document editor for creating and analyzing targeted proteomics experiments.</a> Bioinformatics. 26, 966-968 (2010).</li><li>Proc, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+quantitative+study+of+the+effects+of+chaotropic+agents,+surfactants,+and+solvents+on+the+digestion+efficiency+of+human+plasma+proteins+by+trypsin.">A quantitative study of the effects of chaotropic agents, surfactants, and solvents on the digestion efficiency of human plasma proteins by trypsin.</a> J. Proteome Res. 9, 5422-5437 (2010).</li><li>Berna, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Online+immunoaffinity+liquid+chromatography/tandem+mass+spectrometry+determination+of+a+type+II+collagen+peptide+biomarker+in+rat+urine:+Investigation+of+the+impact+of+collision-induced+dissociation+fluctuation+on+peptide+quantitation.">Online immunoaffinity liquid chromatography/tandem mass spectrometry determination of a type II collagen peptide biomarker in rat urine: Investigation of the impact of collision-induced dissociation fluctuation on peptide quantitation.</a> Anal. Biochem. 356, 235-243 (2006).</li></ol>

No conflicts of interest declared.

The most critical step in the protocol as described is ensuring the beads remain well-mixed during the incubation period. Allowing the beads to settle to the bottom of the well/vial will result in increased variability. It is also important to spin-down any liquid that may remain on the top of the well/vial following the incubation period. Reproducible trypsin digestion is also critical. The procedure for digestion described here has been utilized extensively in our laboratory in conjunction with SISCAPA; however, it is likely alternate methods of digestion could be optimized for a given set of target proteins.15 Elution of free antibody does not interfere with detection of the peptide, likely because the antibody is excluded from the column or elutes later than the peptides, but the antibody can be crosslinked to the Protein G beads prior to affinity enrichment in large experiments or if free antibody becomes a problem. We have also found it useful to place a magnet below the sample plate on the autosampler as well as washing the trap column in the reverse direction of flow (compared to loading) to remove any residual beads or particles. In addition to the magnetic bead approach described, the technique can also be adapted to a column format (i.e. affinity chromatography).12, 16 
Once the user is comfortable with the overall protocol, the technique is also amenable to several modifications that enhance the overall assay.11 First, it is possible to analyze multiple analytes in a single assay by combining antibodies in the enrichment step (i.e. multiplexing). The mass spectrometer is capable of simultaneously analyzing large numbers of analytes. Second, increasing the volume of original sample improves the sensitivity of the assay.

<table border="1">
	<tbody>
 	<tr>
 	<th>Name of the material</th>
 <th>Company</th>
 <th>Catalogue number</th> 
 </tr>
 <tr>
 	<td>1000ul Deep well 96-well plates</td>
 <td>Eppendorf</td>
 <td>951032786</td> 
 </tr>
 <tr>
 	<td>Oasis HLB 1 cc cartridge plate, Sep-pak, 40 mg 96 well plate</td>
 <td>Waters</td>
 <td>186003966</td> 
 </tr>

 <tr>
 	<td>Kingfisher 96 well plate</td>
 <td>Thermo Fisher Scientific</td>
 <td>97002540</td> 
 </tr>
 <tr>
 	<td>Clear 96 well, white wall plate</td>
 <td>Bio-Rad</td>
 <td>HSP9601</td> 
 </tr>
 <tr>
 	<td>Foil cover for 96 well plate</td>
 <td>Excelscientific</td>
 <td>12-169</td>
 </tr>
 <tr>
 	<td>Axymat Sealing mat for 96 well plate</td>
 <td>Axygen</td>
 <td>521-01-151</td>
 </tr>
 <tr>
 	<td>X pierce film</td>
 <td>Sigma-Aldrich</td>
 <td>Z722502</td>
 </tr>
 <tr>
 	<td>Kingfisher magnetic bead processor with PCR magnet head</td>
 <td>Thermo Fisher Scientific</td>
 <td>HSP 9601</td> 
 </tr> 
 </tbody>
</table>
Table 1: Materials
<table border="1">
	<tbody>
 	<tr>
 	<th>Name of the material</th>
 <th>Company</th>
 <th>Catalogue number</th> 
			<th>Comments (optional)</th> 
 </tr>
 <tr>
 	<td>Urea</td>
 <td>Sigma Aldrich</td>
 <td>U6031</td> 
			 <td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Trizma base</td>
 <td>Sigma Aldrich</td>
 <td>T1503</td> 
			<td>&nbsp;</td> 
 </tr>

 <tr>
 	<td>Dithiothreitol (DTT)</td>
 <td>Pierce</td>
 <td>20291</td> 
			<td>"no-weigh dithiothreitol"</td> 
 </tr>
 <tr>
 	<td>Iodoacetamide (IAM)</td>
 <td>Sigma Aldrich</td>
 <td>I1149</td> 
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Trypsin Gold</td>
 <td>Promega</td>
 <td>V5280</td>
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Protein G magnetic beads, 2.8um</td>
 <td>Invitrogen</td>
 <td>10004D</td>
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Phosphate-buffered saline (PBS)</td>
 <td>Thermo Fisher Scientific</td>
 <td>L5401</td>
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>CHAPS </td>
 <td>Thermo Fisher Scientific</td>
 <td>28300 </td> 
			<td>&nbsp;</td> 
 </tr> 
		<tr>
 	<td>Formic acid </td>
 <td>EMD</td>
 <td>11670 </td> 
			<td>&nbsp;</td> 
 </tr> 
		<tr>
 	<td>Acetonitrile </td>
 <td>Thermo Fisher Scientific</td>
 <td>A998-1</td>
			<td>&nbsp;</td> 
 </tr> 
		<tr>
 	<td>Acetic Acid </td>
 <td>Sigma Aldrich</td>
 <td>242853 </td>
			<td>&nbsp;</td> 
 </tr> 
 </tbody>
</table>
Table 2: Reagents
<table border="1">
	<tbody>
 	<tr>
 	<th>Solution</th>
 <th>Comments (optional)</th> 
 </tr>
 <tr>
 	<td>100 mM Tris pH 8</td> 
			 <td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>9 M urea / 30 mM DTT in 100 mM Tris (pH 8)</td>
 <td>must be prepared fresh each time</td> 
 </tr>

 <tr>
 	<td>500 mM IAM in 100 mM Tris (pH 8)</td>
 <td>must be prepared fresh each time</td> 
 </tr>
 <tr>
 	<td>Trypsin 1mg/mL in 100 mM Tris pH 8</td> 
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Trypsin inhibitor 1mg/mL in 100 mM Tris pH 8</td> 
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Stable isotope standard mastermix</td> 
			<td>&nbsp;</td> 
 </tr>
 <tr>
 	<td>Anti-peptide antibody mastermix</td> 
			<td>&nbsp;</td> 
 </tr> 
 </tbody>
</table>
Table 3: Solutions to be prepared

quantification of proteins using peptide immunoaffinity enrichment coupled with mass spectrometry

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in quantitation, are associated with a high cost, long lead time, and are fraught with drawbacks (e.g. heterophilic antibodies, autoantibody interference, 'hook-effect').1 An alternative technique is affinity enrichment of peptides coupled with quantitative mass spectrometry, commonly referred to as SISCAPA (Stable Isotope Standards and Capture by Anti-Peptide Antibodies).2 In this technique, affinity enrichment of peptides with stable isotope dilution and detection by selected/multiple reaction monitoring mass spectrometry (SRM/MRM-MS) provides quantitative measurement of peptides as surrogates for their respective proteins. SRM/MRM-MS is well established for accurate quantitation of small molecules 3, 4 and more recently has been adapted to measure the concentrations of proteins in plasma and cell lysates.5-7 To achieve quantitation of proteins, these larger molecules are digested to component peptides using an enzyme such as trypsin. One or more selected peptides whose sequence is unique to the target protein in that species (i.e. "proteotypic" peptides) are then enriched from the sample using anti-peptide antibodies and measured as quantitative stoichiometric surrogates for protein concentration in the sample. Hence, coupled to stable isotope dilution (SID) methods (i.e. a spiked-in stable isotope labeled peptide standard), SRM/MRM can be used to measure concentrations of proteotypic peptides as surrogates for quantification of proteins in complex biological matrices. The assays have several advantages compared to traditional immunoassays. The reagents are relatively less expensive to generate, the specificity for the analyte is excellent, the assays can be highly multiplexed, enrichment can be performed from neat plasma (no depletion required), and the technique is amenable to a wide array of proteins or modifications of interest.8-13 In this video we demonstrate the basic protocol as adapted to a magnetic bead platform.

Watch this Scientific Journal Video about Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry at JoVE.com

Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry

Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) couples affinity enrichment of peptides with stable isotope dilution mass spectrometry (MRM-MS) to provide quantitative measurement of peptides as surrogates for their respective proteins. Here we describe the protocol using magnetic particles in a partially automated format.

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in ...

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22.7K Views.  Fred Hutchinson Cancer Research Center - FHCRC. Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) couples affinity enrichment of peptides with stable isotope dilution mass spectrometry (MRM-MS) to provide quantitative measurement of peptides as surrogates for their respective proteins. Here we describe the protocol using magnetic particles in a partially automated format.

Video: Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

A Quantitative Assessment of The Yeast Lipidome using Electrospray Ionization Mass Spectrometry

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast Saccharomyces cerevisiae, a genetically and biochemically manipulable unicellular eukaryote with annotated genome and very simple lipidome, is a valuable model for studying biological functions of various lipid species in multicellular eukaryotes2,3,5. S. cerevisiae has 10 major classes of lipids with chain lengths mainly of 16 or 18 carbon atoms and either zero or one degree of unsaturation6,7. Existing methods for lipid identification and quantification - such as high performance liquid chromatography, thin-layer chromatography, fluorescence microscopy, and gas chromatography followed by MS - are well established but have low sensitivity, insufficiently separate various molecular forms of lipids, require lipid derivitization prior to analysis, or can be quite time consuming. Here we present a detailed description of our experimental approach to solve these inherent limitations by using survey-scan ESI/MS for the identification and quantification of the entire complement of lipids in yeast cells. The described method does not require chromatographic separation of complex lipid mixtures recovered from yeast cells, thereby greatly accelerating the process of data acquisition. This method enables lipid identification and quantification at the concentrations as low as g/ml and has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. Lipids extraction from whole yeast cells for using this method of lipid analysis takes two to three hours. It takes only five to ten minutes to run each sample of extracted and dried lipids on a Q-TOF mass spectrometer equipped with a nano-electrospray source.

We describe a new quantitative lipidomics method for identifying numerous lipid species in yeast using survey-scan electrospray ionization mass spectrometry (ESI/MS). This method exceeds currently available methods for lipid identification and quantification in the ability to resolve various molecular forms of lipids, sensitivity, and speed.

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast ...

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an array of dynamic, multi-protein complexes1. To gain a mechanistic understanding of the various cellular processes, it is crucial to determine the structure of such protein complexes, and reveal how their structural organization dictates their function. Many aspects of multi-protein complexes are, however, difficult to characterize, due to their heterogeneous nature, asymmetric structure, and dynamics. Therefore, new approaches are required for the study of the tertiary levels of protein organization.
One of the emerging structural biology tools for analyzing macromolecular complexes is mass spectrometry (MS)2-5. This method yields information on the complex protein composition, subunit stoichiometry, and structural topology. The power of MS derives from its high sensitivity and, as a consequence, low sample requirement, which enables examination of protein complexes expressed at endogenous levels. Another advantage is the speed of analysis, which allows monitoring of reactions in real time. Moreover, the technique can simultaneously measure the characteristics of separate populations co-existing in a mixture. 
Here, we describe a detailed protocol for the application of structural MS to the analysis of large protein assemblies. The procedure begins with the preparation of gold-coated capillaries for nanoflow electrospray ionization (nESI). It then continues with sample preparation, emphasizing the buffer conditions which should be compatible with nESI on the one hand, and enable to maintain complexes intact on the other. We then explain, step-by-step, how to optimize the experimental conditions for high mass measurements and acquire MS and tandem MS spectra. Finally, we chart the data processing and analyses that follow. Rather than attempting to characterize every aspect of protein assemblies, this protocol introduces basic MS procedures, enabling the performance of MS and MS/MS experiments on non-covalent complexes. Overall, our goal is to provide researchers unacquainted with the field of structural MS, with knowledge of the principal experimental tools.

Mass spectrometry has proven to be a valuable tool for analyzing large protein complexes. This method enables insights into the composition, stoichiometry and overall architecture of multi-subunit assemblies. Here, we describe, step-by-step, how to perform a structural mass spectrometry analysis, and characterize macromolecular structures.

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity chromatography 1 or Activity Based Protein Profiling 2. Trifunctional Capture Compounds (CCs, Figure 1A) 3 are the basis for a generic approach, in which the initial equilibrium-driven interaction between a small molecule probe (the selectivity function, here S-adenosyl-L-homocysteine, SAH, Figure 1A) and target proteins is irreversibly fixed upon photo-crosslinking between an independent photo-activable reactivity function (here a phenylazide) of the CC and the surface of the target proteins. The sorting function (here biotin) serves to isolate the CC - protein conjugates from complex biological mixtures with the help of a solid phase (here streptavidin magnetic beads). Two configurations of the experiments are possible: "off-bead" 4 or the presently described "on-bead" configuration (Figure 1B). The selectivity function may be virtually any small molecule of interest (substrates, inhibitors, drug molecules).
S-Adenosyl-L-methionine (SAM, Figure 1A) is probably, second to ATP, the most widely used cofactor in nature 5, 6. It is used as the major methyl group donor in all living organisms with the chemical reaction being catalyzed by SAM-dependent methyltransferases (MTases), which methylate DNA 7, RNA 8, proteins 9, or small molecules 10. Given the crucial role of methylation reactions in diverse physiological scenarios (gene regulation, epigenetics, metabolism), the profiling of MTases can be expected to become of similar importance in functional proteomics as the profiling of kinases. Analytical tools for their profiling, however, have not been available. We recently introduced a CC with SAH as selectivity group to fill this technological gap (Figure 1A).
SAH, the product of SAM after methyl transfer, is a known general MTase product inhibitor 11. For this reason and because the natural cofactor SAM is used by further enzymes transferring other parts of the cofactor or initiating radical reactions as well as because of its chemical instability 12, SAH is an ideal selectivity function for a CC to target MTases. Here, we report the utility of the SAH-CC and CCMS by profiling MTases and other SAH-binding proteins from the strain DH5&alpha; of Escherichia coli (E. coli), one of the best-characterized prokaryotes, which has served as the preferred model organism in countless biochemical, biological, and biotechnological studies. Photo-activated crosslinking enhances yield and sensitivity of the experiment, and the specificity can be readily tested for in competition experiments using an excess of free SAH.

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry CCMS

Capture Compounds are trifunctional small molecules to reduce the complexity of the proteome by functional reversible small molecule-protein interaction followed by photo-crosslinking and purification. Here we use a Capture Compound with S-adenosyl-L-homocysteine-binding as selectivity function to isolate methyltransferases from an Escherichia coli whole cell lysate and identify them by MS.

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity ...

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato3. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas3. Hydrogen peroxide can be detected using a histochemical stain 3'-3' diaminobenzidine (DAB)4. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue4. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins5. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter10. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.

Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

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 ...

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 ...