Name: resin-assisted capture coupled with isobaric tandem mass tag labeling for multiplexed quantification of protein thiol oxidation
Uploaded: 2021-06-21T13:00:00
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 (s...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#8211;Sepharose&#160;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 ...

This protocol describes a method that enables site-specific identification of oxidized cystine residues of proteins. The method allows quantitative and site-specific data of oxidized cystine residues to be generated from various sample types. To begin the assisted capture procedure, wash the precipitated protein pellets twice with acetone by centrifuging at 20, 500 G, in four degrees Celsius for 10 minutes.Then decant the acetone and remove the remaining acetone with a micro pipette. Using a glass serological pipette add three milliliters of fresh ice cold acetone to the tube and mix well by inverting the tube several times. After the second wash, allow the pellets to air dry for one to two minutes without letting them over dry.Add one milliliter of buffer B to the dried protein pellet. To solubilize the protein, sonicate the pellet repeatedly in a bath sonicator with an output of 250 watts for 15 to 30 seconds at a time, along with brief vortexing. Perform the Bicinchoninic acid or BCA assay to measure the protein concentration.To standardize the protein concentrations, transfer 500 micrograms of protein sample to a 0.5 milliliter, 10 kilodaltons centrifugal filter, and volume up the sample to 500 microliters with resuspension buffer. Centrifuge the protein buffer mixture at 14, 000 G in room temperature, until the volume in the centrifugal filter is less than 100 microliters. Collect the sample by inverting the filter in a collection tube.Centrifuge the sample at 1000 G for two minutes and add buffer C to get a final volume of 500 microliters. To reduce the protein thiols, add 20 microliters of 500 millimolar dithiothreitol or DTT, to the sample to get a final concentration of 20 millimolar, and incubate the samples for 30 minutes at 37 degrees Celsius with shaking at 850 RPM. After reduction, centrifuge to the samples into 0.5 milliliter 10 kilodalton centrifugal filters at 14, 000 G at room temperature, until the volume in the centrifugal filter is less than 100 microliters.Add buffer D to make up the volume to 500 microliters and repeat centrifugation. After the fourth centrifugation, collect the samples by inverting the filter in a collection tube to centrifuge at 1000 G for two minutes. Then measure the protein concentration using the BCA assay.Next, place the spin columns on a vacuum manifold, and using a micro pipette transfer 500 microliters of the freshly prepared thiol affinity resin slurry to each column. Apply vacuum for removal of water. Repeat the procedure once to obtain 30 milligrams of resin per column.Wash the resin five times with 500 microliters of ultrapure water, by applying a vacuum to remove the water and then five times with 500 microliters of buffer E.In a new tube, add buffer C to 150 micrograms of the reduced protein sample until the final volume is 120 microliters. Transfer the protein solution to a plugged spin column containing the resin. Place the cap on the column and incubate for two hours at room temperature with shaking at 850 rpm.Wash the resin as described in the manuscript. Replace the plug. To perform on-resin tryptic digestion, add 120 microliters of freshly prepared trypsin solution to the samples, and incubate the column overnight at 37 degrees Celsius, with shaking at 850 RPM.On the next day, wash the resin multiple times as described in the manuscript and replace the plug. Using a micro pipette, add 40 microliters of 100 millimolar Triethylammonium bicarbonate buffer to the washed resin. Then add 70 microliters of the dissolved tandem mass tag or TMT labeling reagent, and incubate for one hour at room temperature with shaking at 850 RPM.Store any remaining TMT reagent at minus 80 degrees Celsius. Wash the resin and replace the plug. Elute the TMT labeled peptides by adding 100 microliters of 100 millimolar ammonium bicarbonate buffer at pH 8.0, containing 20 millimolar DTT, to the column, and incubating the column on a thermal mixer at room temperature for 30 minutes at 850 RPM.After incubation, perform elution by placing the column on a vacuum manifold, applying the vacuum and eluting the samples into a five milliliter micro centrifuge tube. Repeat the step once, then add 100 microliters of 80%acetonitrile with 0.1%trifluoroacetic acid to the column. After incubating the column for 10 minutes at room temperature, elute the sample in the same five milliliter centrifuge tube.Place the eluted samples in a vacuum concentrator until dry. Then store the dry peptides at minus 80 degrees Celsius until further use. The qualitative analysis of peptide samples from RAW 264.7 cells was carried out with SDS-PAGE, wherein samples were compared by determining different levels of oxidation due to treatments.The separated proteins were subsequently probed by western blot to further investigate the oxidation levels of individual proteins. Reported ion intensities of cystine containing peptides were analyzed by liquid chromatography tandem mass spectrometry. The thiol oxidation stoichiometry of iTRAQ, labeled enriched and oxidized peptides at individual cystine site levels were quantified.Carefully remove acetone without disturbing the protein pellet. Ensure that the protein pellet is completely solubilized and quantitated precisely, and that resin is accurately allocated to spin columns. Be sure to resuspend resin during the wash steps.Further evaluation of enriched proteins and peptides may be performed by SDS-PAGE and staining, western blotting and mass spectrometry.

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

Research

JoVE Journal

Biochemistry

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

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 ...

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

A Tandem Liquid Chromatography&#8211;Mass Spectrometry-based Approach for Metabolite Analysis of Staphylococcus aureus

In an effort to thwart bacterial pathogens, hosts often limit the availability of nutrients at the site of infection. This limitation can alter the abundances of key metabolites to which regulatory factors respond, adjusting cellular metabolism. In recent years, a number of proteins and RNA have emerged as important regulators of virulence gene expression. For example, the CodY protein responds to levels of branched-chain amino acids and GTP and is widely conserved in low G+C Gram-positive bacteria. As a global regulator in Staphylococcus aureus, CodY controls the expression of dozens of virulence and metabolic genes. We hypothesize that S. aureus uses CodY, in part, to alter its metabolic state in an effort to adapt to nutrient-limiting conditions potentially encountered in the host environment. This manuscript describes a method for extracting and analyzing metabolites from S. aureus using liquid chromatography coupled with mass spectrometry, a protocol that was developed to test this hypothesis. The method also highlights best practices that will ensure rigor and reproducibility, such as maintaining biological steady state and constant aeration without the use of continuous chemostat cultures. Relative to the USA200 methicillin-susceptible S. aureus isolate UAMS-1 parental strain, the isogenic codY mutant exhibited significant increases in amino acids derived from aspartate (e.g., threonine and isoleucine) and decreases in their precursors (e.g., aspartate and O-acetylhomoserine). These findings correlate well with transcriptional data obtained with RNA-seq analysis: genes in these pathways were up-regulated between 10- and 800-fold in the codY null mutant. Coupling global analyses of the transcriptome and the metabolome can reveal how bacteria alter their metabolism when faced with environmental or nutritional stress, providing potential insight into the physiological changes associated with nutrient depletion experienced during infection. Such discoveries may pave the way for the development of novel anti-infectives and therapeutics.

A Tandem Liquid Chromatography&amp;#8211;Mass Spectrometry-based Approach for Metabolite Analysis of &lt;em&gt;Staphylococcus aureus&lt;/em&gt;

Here we describe a protocol for the extraction of metabolites from Staphylococcus aureus and their subsequent analysis via liquid chromatography and mass spectrometry.

In an effort to thwart bacterial pathogens, hosts often limit the availability of nutrients at the site of infection. This limitation can alter the ...

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) ...

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to precisely maneuver the nucleation and crystal growth towards desired sizes of protein crystals. The controlled crystallization is based on sample investigation with in situ Dynamic Light Scattering (DLS) while all visual changes in the droplet are monitored online with the help of a microscope coupled to a CCD camera, thus enabling a full investigation of the protein droplet during all stages of crystallization. The use of in situ DLS measurements throughout the entire experiment allows a precise identification of the highly supersaturated protein solution transitioning to a new phase &#8211; the formation of crystal nuclei. By identifying the protein nucleation stage, the crystallization can be optimized from large protein crystals to the production of protein microcrystals. The experimental protocol shows an interactive crystallization approach based on precise automated steps such as precipitant addition, water evaporation for inducing high supersaturation, and sample dilution for slowing induced homogeneous nucleation or reversing phase transitions.

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Here we present a protocol for controlled production of protein microcrystals. The process uses an automated device allowing controlled manipulation of several crystallization parameters. The protein crystallization is carried-out by controlled and automated addition of crystallization solutions while monitoring and investigating the radius distribution of particles in the crystallization droplet.

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to ...

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

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

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

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

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...

Detection of Protein S-Acylation using Acyl-Resin Assisted Capture

Protein S-acylation, also referred to as S-palmitoylation, is a reversible post-translational modification of cysteine residues with long-chain fatty acids via a labile thioester bond. S-acylation, which is emerging as a widespread regulatory mechanism, can modulate almost all aspects of the biological activity of proteins, from complex formation to protein trafficking and protein stability. The recent progress in understanding of the biological function of protein S-acylation was achieved largely due to the development of novel biochemical tools allowing robust and sensitive detection of protein S-acylation in a variety of biological samples. Here, we describe acyl resin-assisted capture (Acyl-RAC), a recently developed method based on selective capture of endogenously S-acylated proteins by thiol-reactive Sepharose beads. Compared to existing approaches, Acyl-RAC requires fewer steps and can yield more reliable results when coupled with mass spectrometry for identification of novel S-acylation targets. A major limitation in this technique is the lack of ability to discriminate between fatty acid species attached to cysteines via the same thioester bond.

Acyl-RAC (Acyl-Resin Assisted Capture) is a highly sensitive, reliable and easy to perform method to detect reversible lipid modification of cysteine residues (S-acylation) in a variety of biological samples.

Protein S-acylation, also referred to as S-palmitoylation, is a reversible post-translational modification of cysteine residues with long-chain fatty ...