Name: resin-assisted capture coupled with isobaric tandem mass tag labeling for multiplexed quantification of protein thiol oxidation
Uploaded: 2021-06-21T13:00:00.000+00:00
Duration: 7 min 16 s
Description: Watch this Scientific Journal Video about Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation at JoVE.com

Portions of the work were supported by NIH Grants R01 DK122160, R01 HL139335, and U24 DK112349

All procedures described in the protocol related to animal or human samples/tissues were approved by and followed the institutional guidelines of the human and animal research ethics committee.
1. Sample homogenization/lysis
<ol>
	<li>Frozen tissue samples
	<ol>
		<li>Mince frozen tissue (~30 mg) on a glass microscope slide on dry ice using a prechilled razor blade and forceps. Transfer the minced tissue to a prechilled 5 mL round-bottom polystyrene tube containing 700 &#181;L of buffer A (see Table 1) and incubate on ice for 30 min, protected from light.</li>
		<li>Disrupt the tissue for 30 s or until completely homogenized with a hand-held tissue homogenizer. Place the samples on ice and allow the foam to subside for another 10 min. 
		NOTE: An aluminum baking sheet placed on dry ice provides a stable working surface and platform for the initial processing/mincing of the tissue.</li>
	</ol>
	</li>
	<li>Alternatively, use adherent cell cultures in 100 mm culture dishes as the starting material.
	<ol>
		<li>Keep the cells on ice and use a serological pipette to rinse the cells twice with 10 mL of ice-cold PBS containing 100 mM NEM.</li>
		<li>Lyse the cells by adding 1 mL of cold homogenization/lysis buffer and scraping vigorously with a rigid cell scraper. Transfer the lysate to a 2 mL centrifuge tube using a micropipette. 
		NOTE: Rinsing buffer and lysis buffer may be scaled accordingly to different size culture vessels. Typically, 2-5 million cells are required; however, this varies depending on the lysis efficiency and protein yield for specific cell types. Homogenization buffer may be prepared without NEM for samples being analyzed for total thiols.</li>
	</ol>
	</li>
	<li>Transfer the resulting homogenate (step 1.1.2 or 1.2.2) to a 2 mL centrifuge tube using a micropipette, and centrifuge at full speed (&#8805;16,000 &#215; g) at 4 &#176;C for 10 min.</li>
	<li>Transfer the supernatant (~700 &#181;L or ~1 mL for cell culture) using a micropipette to a 5 mL conical microcentrifuge tube and incubate for 30 min at 55 &#176;C in the dark with shaking at 850 rpm.</li>
	<li>Using a glass serological pipette, add 4 mL of ice-cold acetone to the samples and incubate at -20 &#176;C overnight for precipitation of protein and removal of excess N-ethylmaleimide.</li>
</ol>
2. Resin-assisted capture
<ol>
	<li>Wash the precipitated protein pellets twice with acetone by centrifuging at 20,500 &#215; g at 4 &#176;C for 10 min, decanting the acetone, removing any remaining acetone using a micropipette, and adding 3 mL of fresh, ice-cold acetone using a glass serological pipette. Invert several times to mix. After the second wash, allow the pellets to air-dry for 1-2 min, being careful not to over-dry as resuspension may become difficult.</li>
	<li>Using a micropipette, add 1 mL of buffer B (see Table 1) and solubilize the protein using repeated sonication for 15-30 s at a time using a bath sonicator with an output of 250 W and brief vortexing. Measure the protein concentration using the bicinchoninic acid (BCA) assay according to the manufacturer&#39;s protocol.</li>
	<li>To standardize the protein concentrations across samples for further processing and ensure complete removal of NEM, transfer 500 &#181;g of protein to a 0.5 mL 10 kDa centrifugal filter using a micropipette and adjust to a final volume of 500 &#181;L with resuspension buffer.</li>
	<li>Centrifuge at 14,000 &#215; g at room temperature until the volume in the centrifugal filter is less than 100 &#181;L. Collect the samples by inverting the filter in a collection tube. Centrifuge at 1,000 &#215; g for 2 min and adjust to a final volume of 500 &#181;L using buffer C (see Table 1).</li>
	<li>Reduce the protein thiols by adding 20 &#181;L of 500 mM dithiothreitol (DTT) using a micropipette to a final concentration of 20 mM and incubating the samples for 30 min at 37 &#176;C while shaking at 850 rpm.</li>
	<li>After reduction, transfer the samples using a micropipette to 0.5 mL 10 kDa centrifugal filters and centrifuge for 15 min at 14,000 &#215; g at room temperature or until the volume in the centrifugal filter is less than 100 &#181;L. Add buffer D (see Table 1) to make up the volume in the centrifugal filter to 500 &#181;L.
	<ol>
		<li>Repeat the centrifuging and addition to 500 &#181;L in step 2.6 three times, and after the fourth centrifugation, collect the samples by inverting the filter in a collection tube and centrifuging at 1,000 &#215; g for 2 min.</li>
	</ol>
	</li>
	<li>Measure the protein concentration using the BCA assay according to the manufacturer&#39;s protocol.</li>
	<li>During this buffer exchange, prepare the thiol-affinity resin by weighing the appropriate amount of resin (30 mg/sample) using a microbalance and transferring it to a 50 mL centrifuge tube. Then, using a serological pipette, add water for a final concentration of 30 mg/mL resin and incubate at room temperature for 1 h with agitation for proper hydration of the resin. 
	NOTE: The thiol-affinity resin mentioned above has been discontinued by the manufacturer. A possible replacement for this thiol-affinity resin is commercially available. However, this replacement has a nearly 5-fold less binding capacity (see Supplemental Information). Alternatively, the thiol-affinity resin can be synthesized using 2-(pyridyldithio) ethylamine hydrochloride and N-hydroxysuccinimide-activated resin (see Supplemental Information).
	<ol>
		<li>After hydration of the resin, place the spin columns on a vacuum manifold and transfer 500 &#181;L of the resin slurry using a micropipette to each column. Apply vacuum for removal of water; repeat this step once to obtain a total of 30 mg of resin per column. Alternatively, centrifuge at 1,000 &#215; g for 2 min instead of using the vacuum manifold for this and all the resin washing and elution steps. 
		NOTE: Cutting the end of a 1000 &#181;L pipette tip to increase the bore size helps with the transfer of the resin. It is important to triturate between pipetting to ensure that the resin remains suspended and homogeneous and equal amounts of resin are transferred to each column.</li>
		<li>Wash the resin by adding 500 &#181;L of ultrapure water with a micropipette and applying vacuum for removal of the water; repeat this 5 times. Then, wash the resin 5 times with 500 &#181;L of buffer E (see Table 1). 
		NOTE: Alternatively, centrifugation at 1,000 x g for 2 min may be used in place of a vacuum manifold for all subsequent wash steps. All the proceeding wash steps are performed with a volume of 500 &#181;L. When adding wash buffers to the column, carefully add with enough force to fully resuspend the resin while avoiding splashing and loss of resin; this allows for complete and efficient washing of resin.</li>
	</ol>
	</li>
	<li>Using a micropipette, transfer 150 &#181;g of protein from each reduced sample to a new tube and adjust to a final volume of 120 &#181;L of buffer C (see Table 1). Transfer the protein solution using a micropipette to a plugged spin column containing the resin, place the cap on the column, and incubate for 2 h at room temperature with shaking at 850 rpm.</li>
	<li>Wash the resin five times with 25 mM HEPES, pH 7.0; 8 M urea; followed by five times with 2 M NaCl; followed by five times with 80% acetonitrile (ACN) with 0.1% trifluoroacetic acid (TFA); and finally five times with 25 mM HEPES, pH 7.7, as described in step 2.8.2 and replace the plug. 
	NOTE: Samples may be eluted here for analysis at the protein level (e.g., SDS-polyacrylamide gel electrophoresis (SDS-PAGE), western blot) as described in step 4.1.</li>
</ol>
3. On-resin tryptic digestion and TMT labeling
<ol>
	<li>Prepare enough sequencing-grade modified trypsin solution for 6-8 &#181;g per sample by solubilizing it at a concentration of 0.5 &#181;g/&#181;L in buffer C (see Table 1) so that the final volume allows for at least 120 &#181;L per sample. Using a micropipette, add 120 &#181;L of this trypsin solution to the samples and incubate overnight at 37 &#176;C with shaking at 850 rpm. 
	NOTE: To increase the digestion efficiency, an additional digestion step can be included the next day by removing the trypsin solution and replacing it with fresh solution, and continue the digestion for 2 h.</li>
	<li>Wash the resin five times with 25 mM HEPES, pH 7.0; followed by five times 2 M NaCl; followed by five times with 80% ACN with 0.1% TFA; followed by three times with 25 mM HEPES, pH 7.7. Finally, wash the resin two times with 50 mM triethyl ammonium bicarbonate buffer (TEAB) and replace the plug.</li>
	<li>Prepare TMT labeling reagents by first allowing them to warm to room temperature before spinning down briefly using a centrifuge at 16,000 &#215; g. Add 150 &#181;L of anhydrous ACN to each vial of TMT labeling reagent using a micropipette. Incubate the vials at room temperature on a thermomixer set to 850 rpm for 5 min to solubilize the reagent completely. Briefly vortex and spin down at 16,000 &#215; g to collect the reagent.</li>
	<li>Using a micropipette, add 40 &#181;L of 100 mM TEAB to the washed resin, then add 70 &#181;L of the dissolved TMT reagent and incubate for 1 h at room temperature with shaking at 850 rpm. Store any remaining TMT reagent at -80 &#176;C. 
	NOTE: Take note of the individual TMT labels assigned to each biological sample (Figure 1).</li>
	<li>Wash the resin five times with 80% ACN with 0.1% TFA, three times with 100 mM ammonium bicarbonate buffer (ABC), pH 8.0, and twice with water as previously described and replace the plug.</li>
</ol>
4. Peptide elution
<ol>
	<li>Elute the labeled peptides by adding 100 &#181;L of 20 mM DTT in 100 mM ABC, pH 8.0, to each column using a micropipette and incubate at room temperature for 30 min on a thermomixer set to 850 rpm. 
	NOTE: After the addition of DTT, the resin will clump. The resin can be disrupted with a pipette tip to break up clumps and ensure the complete elution of the peptides.</li>
	<li>After this incubation, place the column on a vacuum manifold intended for solid-phase extraction (SPE), apply vacuum, and elute the samples into a 5 mL microcentrifuge tube. Repeat this step once.</li>
	<li>Finally, add 100 &#181;L of 80% ACN with 0.1% TFA, incubate for 10 min at room temperature, and elute into the same 5 mL centrifuge tube. Collect all the fractions in the same 5 mL microcentrifuge tube. 
	NOTE: To prevent sample loss, low binding tubes must be used for elution, and volumes must be kept at or below a volume of 4.0 mL&#160;for a single 5 mL tube.</li>
	<li>Place the eluted samples in a vacuum concentrator until dry. Store the dry peptides at -80 &#176;C and resuspend them later. 
	&#8203;NOTE: Samples may also be eluted separately, and an aliquot may be removed and analyzed by SDS-PAGE for analysis at the peptide level before combining the samples.</li>
</ol>
5. Peptide alkylation and desalting/clean-up
<ol>
	<li>Resuspend the dried peptides by adding a small volume of 100 mM ABC buffer, pH 8.0 (no greater than 500 &#181;L), using a micropipette. Use repeated sonication for 15-30 s at a time using a bath sonicator with an output of 250 W and vortex to solubilize and transfer to a 2 mL tube. 
	NOTE: The volume of 100 mM ABC, pH 8.0 to be added is based on the volume needed to resuspend the DTT at a molarity of 150 mM. Users will need to determine the amount of DTT present in their sample based on what was added originally in step 4.1.</li>
	<li>Add enough concentrated stock solution (600 mM) of iodoacetamide (IAA) dissolved in ABC using a micropipette to achieve a 1:4 molar ratio of DTT:IAA and incubate the samples at RT for 1 h with shaking at 850 rpm.</li>
	<li>Acidify the samples to pH &#60; 3 by adding concentrated TFA (10%) using a micropipette and perform sample desalting using reverse-phase clean-up according to the manufacturer&#39;s instructions.</li>
	<li>Place the clean peptides in a vacuum concentrator until dry. Store the dry peptides at -80 &#176;C until further analysis.</li>
</ol>
6. Liquid chromatography-tandem mass spectrometry
<ol>
	<li>Resuspend the dried peptides by repeated sonication for 15-30 s at a time using a bath sonicator with an output of 250 W and vortexing in 20-40 &#181;L of water containing 3% ACN. Determine the peptide concentration by performing a BCA assay according to the manufacturer&#39;s protocol.</li>
	<li>Separate the samples by reversed-phase LC and MS/MS as previously described6 and record the MS1 spectra over the m/z range of 400-2000. Ensure high-energy collisional dissociation (HCD) is utilized to obtain reporter ion intensity information for the analysis of isobarically labeled peptides. See the methods sections of previous reports for more details about instrument run conditions27,30 and the analysis of MS data27,31. 
	NOTE: Different LC-MS/MS systems or settings can be used to analyze the peptide samples. The coverage and sensitivity of peptide identification will depend on the particular system and settings used.</li>
</ol>

The authors declare no conflicts of interest, financial or otherwise.

Resin-assisted capture has been utilized across a variety of sample types and biological systems for the investigation of oxidative modifications of cysteine residues25,29,30. This method allows for the evaluation of samples at multiple levels and readouts, including proteins and peptides using SDS-PAGE and western blot analysis, as well as individual cysteine sites using mass spectrometry. Regardless of the sample type or the f...

Under homeostatic conditions, cells generate reactive oxygen, nitrogen, or sulfur species that help to facilitate processes, such as metabolism and signaling1,2,3, extending to both prokaryotes and eukaryotes. Physiological levels of these reactive species are necessary for proper cellular function, also known as &#39;eustress&#39;1,4. In contrast, an increase in oxidants that leads to an imbalance between oxidants and antioxidants can cause oxidative stress, or &#39;distress&#39;1, which leads to cellular damage. Oxidants transduce signals to biological pathways by modifying different biomolecules, including protein, DNA, RNA, and lipids. In particular, cysteine residues of proteins are highly reactive sites prone to oxidation due to the thiol group on cysteine, which is reactive towards different types of oxidants5. This gives rise to a diverse range of reversible redox-based posttranslational modifications (PTMs) for cysteine, including nitrosylation (SNO), glutathionylation (SSG), sulfenylation (SOH), persulfidation (SSH), polysulfidation (SSnH), acylation, and disulfides. Irreversible forms of cysteine oxidation include sulfinylation (SO2H) and sulfonylation (SO3H).
Reversible oxidative modifications of cysteine residues may serve protective roles preventing further irreversible oxidation or serve as signaling molecules for downstream cellular pathways6,7. The reversibility of some thiol redox PTMs allows cysteine sites to function as &#34;redox switches&#34;8,9, wherein changes in the redox state of these sites alter protein function to regulate their role in transient processes. The modulatory effects of redox PTMs10 have been observed in many aspects of protein function11, including catalysis12, protein-protein interactions13, conformation change14, metal ion coordination15, or pharmacological inhibitor binding16. Additionally, redox PTMs are involved in cysteine sites of proteins that regulate pathways such as transcription17, translation18, or metabolism19. Given the impact that redox PTMs have on protein function and biological processes, it is important to quantify the extent of oxidation that a cysteine site undergoes in response to a perturbation of the redox state.
The identification of cysteine sites with altered redox states is focused on the comparison of the oxidation state at the site-specific level between normal and perturbed conditions. Fold change measurements are often utilized to determine what sites are significantly altered as this helps users interpret what cysteine sites may be physiologically significant to the study. Alternatively, stoichiometric measurements of reversible thiol oxidation across a specific sample type give a general picture of the physiological state with respect to cellular oxidation, an important measurement that is often overlooked and underutilized. Modification stoichiometry is based on quantifying the percentage of modified thiol as a ratio to total protein thiol (modified and unmodified)20,21. As a result, stoichiometric measurements offer a more precise measurement than fold change, especially when using mass spectrometry. The significance of the increase in oxidation can be more readily ascertained by using stoichiometry to determine the PTM occupancy of a particular cysteine site. For example, a 3-fold increase in thiol oxidation could result from a transition of as little as 1% to 3% or as big as 30% to 90%. A 3-fold increase in oxidation for a site that is only at 1% occupancy may have little impact on a protein&#39;s function; however, a 3-fold increase for a site with 30% occupancy at resting state may be more substantially affected. Stoichiometric measurements, when performed between total oxidized thiols and specific oxidative modifications, including protein glutathionylation (SSG) and nitrosylation (SNO), can reveal ratios and quantitative information with respect to specific modification types.
Because reversible thiol oxidation is typically a low-abundance posttranslational modification, multiple approaches have been developed for the enrichment of proteins containing these modifications out of biological samples. An early approach devised by Jaffrey and others, named the biotin switch technique (BST)22, involves multiple steps wherein unmodified thiols are blocked through alkylation, reversibly modified thiols are reduced to nascent free thiols, nascent free thiols are labeled with biotin, and the labeled proteins are enriched by streptavidin affinity pulldown. This technique has been used to profile SNO and SSG in many studies and can be adapted to probe for other forms of reversible thiol oxidation23,24. While BST has been utilized to probe for different forms of reversible thiol oxidation, one concern with this approach is that enrichment is impacted by the non-specific binding of unbiotinylated proteins to streptavidin. An alternate approach developed in our&#160;laboratory, named resin-assisted capture (RAC)25,26 (Figure 1), circumvents the issue of enrichment of thiol groups via the biotin-streptavidin system.
Following the reduction of reversibly oxidized thiols, proteins with nascent free thiols are enriched by the thiol-affinity&#160;resin, which covalently captures free thiol groups, allowing for more specific enrichment of cysteine-containing proteins than BST. Coupling RAC with the multiplexing power of the recent advances in isobaric labeling and mass spectrometry creates a robust and sensitive workflow for the enrichment, identification, and quantification of reversibly oxidized cysteine residues at the proteome-wide level. Recent advances in mass spectrometry have enabled much deeper profiling of the thiol redox proteome, increasing the understanding of both the cause and effect of protein thiol oxidation27. The information gained from site-specific quantitative data allows for further studies of the mechanistic impacts and downstream effects of reversible oxidative modifications28. Utilizing this workflow has provided insight into the physiological impacts of reversible cysteine oxidation with respect to normal physiological events such as aging, wherein levels of SSG differed with respect to age. The aging effects on SSG were partially reversed using SS-31 (elamipretide), a novel peptide that enhances mitochondrial function and reduces SSG levels in aged mice, causing them to have an SSG profile more similar to young mice29.
Pathophysiological conditions attributed to nanoparticle exposure have been shown to involve SSG in a mouse macrophage model. Using RAC coupled with mass spectrometry, the authors showed that SSG levels were directly correlated to the degree of oxidative stress and impairment of macrophage phagocytic function. The data also revealed pathway-specific differences in response to different engineered nanomaterials that induce different degrees of oxidative stress30. The method has also proven its utility in prokaryotic species, where it was applied to study the effects of diurnal cycles in photosynthetic cyanobacteria with respect to thiol oxidation. Broad changes in thiol oxidation across several key biological processes were observed, including electron transport, carbon fixation, and glycolysis. Furthermore, through orthogonal validation, several key functional sites were confirmed to be modified, suggesting regulatory roles of these oxidative modifications6.
Herein, we describe the details of a standardized workflow (Figure 1), demonstrating the utility of the RAC approach for the enrichment of total oxidized cysteine thiols of proteins and their subsequent labeling and stoichiometric quantification. This workflow has been implemented in studies of the redox state in different sample types, including cell cultures27,30 and whole tissues (e.g., skeletal muscle, heart, lung)29,31,32,33. While not included here, the RAC protocol is also easily adapted for the investigation of specific forms of reversible redox modifications, including SSG, SNO, and S-acylation, as previously mentioned25,29,34.

<table><tbody><tr><td>2-(Pyridyldithio)ethylamine hydrochloride</td><td>Med Chem Express</td><td>HY-101794</td><td>Reagent for in-house resin synthesis</td></tr><tr><td>2.0 mL LoBind centrifuge tubes</td><td>Eppendorf</td><td>22431048</td><td></td></tr><tr><td>5.0 mL LoBind centrifuge tubes</td><td>Eppendorf</td><td>30108310</td><td></td></tr><tr><td>5.0 mL round bottom tubes</td><td>Falcon</td><td>352054</td><td></td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>A949-1</td><td></td></tr><tr><td>Acetonitrile</td><td>Sigma Aldrich</td><td>34998</td><td></td></tr><tr><td>Activated Thiol&amp;ndash;Sepharose&amp;nbsp;4B</td><td>Sigma Aldrich</td><td>T8512</td><td>Potential replacement for thiol-affinity resin</td></tr><tr><td>Amicon Ultra 0.5 mL centrifugal filter</td><td>Millipore Sigma</td><td>UFC5010BK</td><td></td></tr><tr><td>Ammonium bicarbonate</td><td>Sigma Aldrich</td><td>09830</td><td></td></tr><tr><td>Bicinchonicic acid (BCA)</td><td>Thermo Scientific</td><td>23227</td><td>Protein Assay Reagent</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5810R</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5415R</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>Thermo Scientific</td><td>20291</td><td></td></tr><tr><td>EDTA</td><td>Sigma Aldrich</td><td>E5134</td><td></td></tr><tr><td>HEPES buffer</td><td>Sigma Aldrich</td><td>H4034</td><td></td></tr><tr><td>Homogenizer</td><td>BioSpec Products</td><td>985370</td><td></td></tr><tr><td>Iodoacetimide (IAA)</td><td>Sigma Aldrich</td><td>I1149</td><td></td></tr><tr><td>N-ethylmaleimide</td><td>Sigma Aldrich</td><td>4259</td><td></td></tr><tr><td>NHS-Activated Sepharose 4 Fast Flow</td><td>Cytiva</td><td>17-0906-01</td><td>Reagent for in-house resin synthesis</td></tr><tr><td>QIAvac 24 Plus vacuum manifold</td><td>Qiagen</td><td>19413</td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma Aldrich</td><td>S3014</td><td></td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Sigma Aldrich</td><td>L6026</td><td></td></tr><tr><td>Sonicator</td><td>Branson</td><td>1510R-MT</td><td></td></tr><tr><td>Spin columns</td><td>Thermo Scientific</td><td>69705</td><td></td></tr><tr><td>Strata C18-E reverse phase columns</td><td>Phenomenex</td><td>8B-S001-DAK</td><td>Peptide desalting</td></tr><tr><td>Thermomixer</td><td>Eppendorf</td><td>5355</td><td></td></tr><tr><td>Thiopropyl Sepharose 6B</td><td>GE Healthcare</td><td>17-0420-01</td><td>Thiol-affinity resin; *Production of Thiopropyl Sepharose 6B resin has been discontinued by the manufacturer (see protocol for details).</td></tr><tr><td>TMT isobaric labels (16 plex)</td><td>Thermo Scientific</td><td>A44522</td><td>Peptide labeling reagent; available in multiple formats</td></tr><tr><td>Triethylammonium bicarbonate buffer (TEAB)</td><td>Sigma Aldrich</td><td>T7408</td><td></td></tr><tr><td>Trifluoroacetic acid (TFA)</td><td>Sigma Aldrich</td><td>T6508</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T8787</td><td></td></tr><tr><td>Trypsin</td><td>Promega</td><td>V5820</td><td></td></tr><tr><td>Urea</td><td>Sigma Aldrich</td><td>U5378</td><td></td></tr><tr><td>Vacufuge Plus speedvac</td><td>Eppendorf</td><td>22820001</td><td>vacuum concentrator</td></tr><tr><td>Vortex mixer</td><td>Scientific Industries</td><td>SI-0236</td><td></td></tr></tbody></table>

resin-assisted capture coupled with isobaric tandem mass tag labeling for multiplexed quantification of protein thiol oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Completion of the protocol will result in highly specific enrichment of formerly oxidized cysteine-containing peptides, often with &#62;95% specificity27,35,36. However, several key steps of the protocol require special attention, e.g., the initial blocking of free thiols prior to sample lysis/homogenization, which prohibits artificial oxidation and non-specific enrichment of artificially oxidized thiols25</sup...

Watch this Scientific Journal Video about Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation at JoVE.com

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

resin-assisted-capture-coupled-with-isobaric-tandem-mass-tag-labeling

Research

JoVE Journal

Biochemistry

1.7K Views.  Pacific Northwest National Laboratory. Dit protocol beschrijft een methode die locatiespecifieke identificatie van geoxideerde cystineresiduen van eiwitten mogelijk maakt. De methode maakt het mogelijk kwantitatieve en locatiespecifieke gegevens van geoxideerde cystineresiduen te genereren uit verschillende monstertypen. Om de geassisteerde vangstprocedure te starten, wast u de neergeslagen eiwitkorrels tweemaal met aceton door centrifugeren op 20.500 G, in vier graden Celsius gedurende 10 minuten.Decanteer vervolgens de ac...

Video: Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Quantitative Proteomics Using Reductive Dimethylation for Stable Isotope Labeling

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a method to accurately quantify protein expression differences between samples using mass spectrometry. ReDi labeling is performed using either regular (light) or deuterated (heavy) forms of formaldehyde and sodium cyanoborohydride to add two methyl groups to each free amine. Here we demonstrate a robust protocol for ReDi labeling and quantitative comparison of complex protein mixtures. Protein samples for comparison are digested into peptides, labeled to carry either light or heavy methyl tags, mixed, and co-analyzed by LC-MS/MS. Relative protein abundances are quantified by comparing the ion chromatogram peak areas of heavy and light labeled versions of the constituent peptide extracted from the full MS spectra. The method described here includes sample preparation by reversed-phase solid phase extraction, on-column ReDi labeling of peptides, peptide fractionation by basic pH reversed-phase (BPRP) chromatography, and StageTip peptide purification. We discuss advantages and limitations of ReDi labeling with respect to other methods for stable isotope incorporation. We highlight novel applications using ReDi labeling as a fast, inexpensive, and accurate method to compare protein abundances in nearly any type of sample.

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a rapid, inexpensive strategy for accurate mass spectrometry-based quantitative proteomics. Here we demonstrate a robust method for preparation and analysis of protein mixtures using the ReDi approach that can be applied to nearly any sample type.

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a method to accurately quantify protein expression differences between ...

Chemistry

Enhanced Sample Multiplexing of Tissues Using Combined Precursor Isotopic Labeling and Isobaric Tagging (cPILOT)

There is an increasing demand to analyze many biological samples for disease understanding and biomarker discovery. Quantitative proteomics strategies that allow simultaneous measurement of multiple samples have become widespread and greatly reduce experimental costs and times. Our laboratory developed a technique called combined precursor isotopic labeling and isobaric tagging (cPILOT), which enhances sample multiplexing of traditional isotopic labeling or isobaric tagging approaches. Global cPILOT can be applied to samples originating from cells, tissues, bodily fluids, or whole organisms and gives information on relative protein abundances across different sample conditions. cPILOT works by 1) using low pH buffer conditions to selectively dimethylate peptide N-termini and 2) using high pH buffer conditions to label primary amines of lysine residues with commercially-available isobaric reagents (see Table of Materials/Reagents). The degree of sample multiplexing available is dependent on the number of precursor labels used and the isobaric tagging reagent. Here, we present a 12-plex analysis using light and heavy dimethylation combined with six-plex isobaric reagents to analyze 12 samples from mouse tissues in a single analysis. Enhanced multiplexing is helpful for reducing experimental time and cost and more importantly, allowing comparison across many sample conditions (biological replicates, disease stage, drug treatments, genotypes, or longitudinal time-points) with less experimental bias and error. In this work, the global cPILOT approach is used to analyze brain, heart, and liver tissues across biological replicates from an Alzheimer&#39;s disease mouse model and wild-type controls. Global cPILOT can be applied to study other biological processes and adapted to increase sample multiplexing to greater than 20 samples.

Enhanced Sample Multiplexing of Tissues Using Combined Precursor Isotopic Labeling and Isobaric Tagging cPILOT

Combined precursor isotopic labeling and isobaric tagging (cPILOT) is a quantitative proteomics strategy that enhances sample multiplexing capabilities of isobaric tags. This protocol describes the application of cPILOT to tissues from an Alzheimer's disease mouse model and wild-type controls.

There is an increasing demand to analyze many biological samples for disease understanding and biomarker discovery. Quantitative proteomics strategies ...

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

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

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

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

TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view of the impact of a disease, treatment, or other condition on the proteome as a whole. This report provides a detailed protocol for the extraction, quantification, precipitation, digestion, labeling, and subsequent data analysis of protein samples. Our optimized TMT labeling protocol requires a lower tag-label concentration and achieves consistently reliable data. We have used this protocol to evaluate protein expression profiles in a variety of mouse tissues (i.e., heart, skeletal muscle, and brain) as well as cells cultured in vitro. In addition, we demonstrate how to evaluate thousands of proteins from the resulting dataset.

We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view ...

Automated Sample Multiplexing by using Combined Precursor Isotopic Labeling and Isobaric Tagging (cPILOT)

We have introduced a high throughput quantitative proteomics workflow, combined precursor isotopic labeling and isobaric tagging (cPILOT) capable of multiplexing up to 22 or 24 samples with tandem mass tags or isobaric N,N-dimethyl leucine isobaric tags, respectively, in a single experiment. This enhanced sample multiplexing considerably reduces mass spectrometry acquisition times and increases the utility of the expensive commercial isobaric reagents. However, the manual process of sample handling and pipetting steps in the strategy can be labor intensive, time consuming, and introduce sample loss and quantitative error. These limitations can be overcome through the incorporation of automation. Here we transferred the manual cPILOT protocol to an automated liquid handling device that can prepare large sample numbers (i.e., 96 samples) in parallel. Overall, automation increases feasibility and reproducibility of cPILOT and allows for broad usage by other researchers with comparable automation devices.

Automated Sample Multiplexing by using Combined Precursor Isotopic Labeling and Isobaric Tagging cPILOT

Combined precursor isotopic labeling and isobaric tagging (cPILOT) is an enhanced sample multiplexing strategy that is capable of increasing the number of samples that can be analyzed simultaneously with available isobaric tags. Incorporation of a robotic platform has greatly increased experimental throughput, reproducibility, and quantitative accuracy.

We have introduced a high throughput quantitative proteomics workflow, combined precursor isotopic labeling and isobaric tagging (cPILOT) capable of ...

A Mass Spectrometry-Based Approach to Identify Phosphoprotein Phosphatases and their Interactors

Most cellular processes are regulated by dynamic protein phosphorylation. More than three-quarters of proteins are phosphorylated, and phosphoprotein phosphatases (PPPs) coordinate over 90% of all cellular serine/threonine dephosphorylation. Deregulation of protein phosphorylation has been implicated in the pathophysiology of various diseases, including cancer and neurodegeneration. Despite their widespread activity, the molecular mechanisms controlling PPPs and those controlled by PPPs are poorly characterized. Here, a proteomic approach termed phosphatase inhibitor beads and mass spectrometry (PIB-MS) is described to identify and quantify PPPs, their posttranslational modifications, and their interactors in as little as 12 h using any cell line or tissue. PIB-MS utilizes a non-selective PPP inhibitor, microcystin-LR (MCLR), immobilized on sepharose beads to capture and enrich endogenous PPPs and their associated proteins (termed the PPPome). This method does not require the exogenous expression of tagged versions of PPPs or the use of specific antibodies. PIB-MS offers an innovative way to study the evolutionarily conserved PPPs and expand our current understanding of dephosphorylation signaling.

Here, we present a protocol for the enrichment of endogenous phosphoprotein phosphatases and their interacting proteins from cells and tissues and their identification and quantification by mass spectrometry-based proteomics.

Most cellular processes are regulated by dynamic protein phosphorylation. More than three-quarters of proteins are phosphorylated, and phosphoprotein ...

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 ...

Biology

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 ...

Simultaneous Affinity Enrichment of Two Post-Translational Modifications for Quantification and Site Localization

Studying multiple post-translational modifications (PTMs) of proteins is a crucial step to understand PTM crosstalk and gain more holistic insights into protein function. Despite the importance of multi-PTM enrichment studies, few studies investigate more than one PTM at a time, due partially to the expenses, time, and large protein quantities required to perform multiple global proteomic analysis of PTMs. The &#34;one-pot&#34; affinity enrichment detailed in this protocol overcomes these barriers by permitting the simultaneous identification and quantification of peptides with lysine residues containing acetylation and succinylation PTMs with low amounts of sample input. The protocol involves preparation of protein lysate from mouse livers of SIRT5 knockout mice, performance of trypsin digestion, enrichment for PTMs, and performance of mass spectrometric analysis using a data-independent acquisition (DIA) workflow. Because this workflow allows for the enrichment of two PTMs from the same sample simultaneously, it provides a practical tool to study PTM crosstalk without requiring large amounts of samples, and it greatly reduces the time required for sample preparation, data acquisition, and analysis. The DIA component of the workflow provides comprehensive PTM-specific information. This is particularly important when studying PTM site localization, as DIA provides comprehensive sets of fragment ions that can be computationally deciphered to differentiate between different PTM localization isoforms.

This workflow describes the performance of time- and cost-efficient enrichment of multiple protein post-translational modifications (PTMs) simultaneously for quantitative global proteomic analysis. The protocol utilizes peptide-level PTM enrichment with multiple conjugated antibodies, followed by data-independent acquisition mass spectrometry analysis to gain biological insights into PTM crosstalk.

Studying multiple post-translational modifications (PTMs) of proteins is a crucial step to understand PTM crosstalk and gain more holistic insights into ...

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

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

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

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

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Samenvatting

Meer video's verkennen

Hoofdstukken in deze video