Name: deep proteome profiling by isobaric labeling, extensive liquid chromatography, mass spectrometry, and software-assisted quantification
Uploaded: 2017-11-15T13:00:00
Description: Watch this Scientific Journal Video about Deep Proteome Profiling by Isobaric Labeling, Extensive Liquid Chromatography, Mass Spectrometry, and Software-assisted Quantification at JoVE.com

The authors thank all other lab and facility members for helpful discussion. This work was partially supported by NI H grants R01GM114260, R01AG047928, R01AG053987, and ALSAC. The MS analysis was performed in the St. Jude Children&#39;s Research Hospital Proteomics Facility, partially supported by NIH Cancer Center Support grant P30CA021765. The authors thank Nisha Badders for help with editing the manuscript.

CAUTION: Please consult all relevant safety data sheets (i.e., MSDS) before use. Please use all appropriate safety practices when performing this protocol.
NOTE: A TMT 10-plex isobaric label reagent set is used in this protocol for the proteome quantitation of 10 samples.
1. Preparation of Cells/Tissues
NOTE: It is critical to collect samples in minimal time at low temperature to keep proteins in their original biological state.</p...

<ol>
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We describe a high-throughput protocol for the quantitation of proteins with a 10-plex isobaric labeling strategy, which has been implemented successfully in several publications12,13,14,32. In this protocol, we can analyze up to 10 different biological protein samples in 1 experiment. We can routinely identify and quantitate well over 10,000 proteins with high confidence. Although isobaric lab...

Advances in next-generation sequencing technology have led to a new landscape for studying biological systems and human disease. This has permitted a large number of measurements of the genome, transcriptome, proteome, metabolome, and other molecular systems to become tangible. Mass spectrometry (MS) is one of the most sensitive methods in analytical chemistry, and its application in proteomics has rapidly expanded after the sequencing of the human genome. In the proteomics field, the past few years have yielded major technical advances in MS-based quantitative analyses, including isobaric labeling and multiplexing capability combined with extensive liquid chromatography, in addition to instrumentation advances, allowing for faster, more accurate measurements with less sample material required. Quantitative proteomics have become a mainstream approach for profiling tens of thousands of proteins and posttranslational modifications in highly complex biological samples1,2,3,4,5,6.
Multiplexed isobaric labeling methods such as isobaric tag for relative and absolute quantitation (i.e., iTRAQ) and tandem mass tag (TMT) MS have greatly improved sample throughput and increased the number of samples that can be analyzed in a single experiment1,6,7,8. Along with other MS-based quantitation methods, such as label-free quantitation and stable isotope labeling with amino acids in cell culture (i.e., SILAC), the potential of these techniques in the proteomics field is considerable9,10,11. For example, the TMT method permits 10 protein samples to be analyzed together in 1 experiment by using 10-plex reagents. These structurally identical TMT tags have the same overall mass, but heavy isotopes are differentially distributed on carbon or nitrogen atoms, resulting in a unique reporter ion during MS/MS fragmentation of each tag, thereby enabling relative quantitation between the 10 samples. The TMT strategy is routinely applied to study biological pathways, disease progression, and cellular processes12,13,14.
Substantial technical improvements have enhanced liquid chromatography (LC) &#8211;MS/MS systems, both in terms of LC separations and MS parameters, to maximize protein identification without sacrificing quantitation accuracy. First-dimension separation of peptides by a separation technique with high orthogonality to the second dimension is critical in this type of shotgun proteomics method to achieve maximum results20. High-pH reversed-phase liquid chromatography (RPLC) provides better performance than does conventional strong cation-exchange chromatography20. When high-pH RPLC is combined with a second dimension of low-pH RPLC, both analytical dynamic range and protein coverage are improved, resulting in the ability to identify the bulk of expressed proteins when performing whole-proteome analyses15,16,17,18. Other technical advances include small C18 particles (1.9 &#181;m) and extended long column (~1 m)19. Furthermore, other notable improvements include new versions of mass spectrometers with rapid scan rates, improved sensitivity and resolution20, and sophisticated bioinformatics pipelines for MS data mining21.
Here, we describe a detailed protocol that incorporates the most recent methodologies with modifications to improve both sensitivity and throughput, while focusing on quality control mechanisms throughout the experiment. The protocol includes protein extraction and digestion, TMT 10-plex labeling, basic pH and acid pH RPLC fractionation, high-resolution MS detection, and MS data processing (Figure 1). Moreover, we implement several quality control steps for troubleshooting and evaluating experimental variation. This detailed protocol is intended to help researchers new to the field routinely identify and accurately quantitate thousands of proteins from a lysate or tissue.

<table>
	<tbody>
		<tr>
			<td>1220 LC system</td>
			<td>Agilent</td>
			<td>G4288B</td>
			<td></td>
		</tr>
		<tr>
			<td>50% Hydroxylamine</td>
			<td>Thermo Scientific</td>
			<td>90115</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetonitrile</td>
			<td>Burdick &#38; Jackson</td>
			<td>AH015-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Bullet Blender</td>
			<td>Next Advance</td>
			<td>BB24-AU</td>
			<td></td>
		</tr>
		<tr>
			<td>Butterfly Portfolio Heater</td>
			<td>Phoenix S&#38;T</td>
			<td>PST-BPH-20</td>
			<td></td>
		</tr>
		<tr>
			<td>C18 tips</td>
			<td>Harvard Apparatus</td>
			<td>74-4607</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol (DTT)</td>
			<td>Sigma</td>
			<td>D5545</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma</td>
			<td>41648</td>
			<td></td>
		</tr>
		<tr>
			<td>Formic acid</td>
			<td>Sigma</td>
			<td>94318</td>
			<td></td>
		</tr>
		<tr>
			<td>Fraction Collector</td>
			<td>Gilson</td>
			<td>FC203B</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Beads</td>
			<td>Next Advance</td>
			<td>GB05</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>Iodoacetamide (IAA)</td>
			<td>Sigma</td>
			<td>I6125</td>
			<td></td>
		</tr>
		<tr>
			<td>Lys-C</td>
			<td>Wako</td>
			<td>125-05061</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Burdick &#38; Jackson</td>
			<td>AH230-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Pierce BCA Protein Assay kit</td>
			<td>Thermo Scientific</td>
			<td>23225</td>
			<td></td>
		</tr>
		<tr>
			<td>Mass Spectrometer</td>
			<td>Thermo Scientific</td>
			<td>Q Exactive HF</td>
			<td></td>
		</tr>
		<tr>
			<td>nanoflow UPLC</td>
			<td>Thermo Scientific</td>
			<td>Ultimate 3000</td>
			<td></td>
		</tr>
		<tr>
			<td>ReproSil-Pur C18 resin, 1.9um</td>
			<td>Dr. Maisch GmbH</td>
			<td>r119.aq.0003</td>
			<td></td>
		</tr>
		<tr>
			<td>Self Pck Columns</td>
			<td>New Objective</td>
			<td>PF360-75-15-N-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>Sigma</td>
			<td>30970</td>
			<td></td>
		</tr>
		<tr>
			<td>Speedva</td>
			<td>Thermo Scientific</td>
			<td>SPD11V</td>
			<td></td>
		</tr>
		<tr>
			<td>TMT 10plex Isobaric label reagent</td>
			<td>Thermo Scientific</td>
			<td>90110</td>
			<td></td>
		</tr>
		<tr>
			<td>Trifluoroacetic acid (TFA)</td>
			<td>Applied Biosystems</td>
			<td>400003</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Promega</td>
			<td>V511C</td>
			<td></td>
		</tr>
		<tr>
			<td>Urea</td>
			<td>Sigma</td>
			<td>U5378</td>
			<td></td>
		</tr>
		<tr>
			<td>Xbridge Column C18 column</td>
			<td>Waters</td>
			<td>186003943</td>
			<td></td>
		</tr>
		<tr>
			<td>Ziptips C18</td>
			<td>Millipore</td>
			<td>ZTC18S096</td>
			<td></td>
		</tr>
		<tr>
			<td>SepPak 1cc 50mg</td>
			<td>Waters</td>
			<td>WAT054960</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 used a previously described cross-species peptide mix to systematically analyze the effect of ratio compression in 3 major protocol steps, including pre-MS fractionation, MS settings, and post-MS correction23. The pre-MS fractionation was evaluated and optimized by using a combination of basic pH RPLC and acidic pH RPLC. For post-MS analysis, only species-specific peptides were considered. We used this interference model to examine a number of parameters in LC/L...

Watch this Scientific Journal Video about Deep Proteome Profiling by Isobaric Labeling, Extensive Liquid Chromatography, Mass Spectrometry, and Software-assisted Quantification at JoVE.com

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

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) ...

The overall goal of this experiment is to identify and accurately quantitate over 10, 000 proteins from cell or tissue lysate across different drug treatments, time courses, or biological conditions. This method can help answer key questions in the field of medicine or biology such as what proteins are changing in response to a drug or other biological stimulus. The main advantage of this technique is that it has the ability to accurately quantitate over 70%of the expressed proteome over 10 different experimental conditions.The implication of this technology can be extended to the diagnosis of human diseases through potential biomarker discovery. Visual demonstration of this method is critical as optimal cell lysis and fluid estimation play a key role in downstream digestion, labeling, and separation procedures. To begin this protocol, obtain cultured cells of interest.Wash the cells twice using 10 milliliters of PBS for each wash. Scrape and collect the washed cells in a 1.5 milliliter tube containing one milliliter of PBS. Centrifuge at 600 times g and four degrees Celsius for five minutes.Remove the supernatant and store at minus 80 degrees Celsius until ready to lyse the cells. Collect and weigh the desired tissues quickly after dissection. After this, immediately store them in liquid nitrogen and store at minus 80 degrees Celsius.Prepare the lysis buffer on the day of the experiment as outlined in the text protocol. Add lysis buffer to the frozen sample such that the buffer to sample ratio is 10 to one and the final protein concentration is between five and 10 milligrams per milliliter. Next, add glass beads and use a blender to lyse the samples at four degrees Celsius and speed eight by blending for 30 seconds and then resting for five seconds, repeating this five times or until the samples are homogenized.Measure the protein concentration using a standard protein quantitation assay or a Coomassie-stained SDS polyacrylamide gel with BSA as a standard. After this, add 100%acetonitrile such that the final concentration is 10%and Lys-C protease is at an enzyme to substrate ratio of one to 100. Incubate at room temperature for two hours to allow protein digestion to occur.Then add DTT such that the final concentration is one millimolar. Incubate at room temperature for one hour. Next, add 50 millimolar HEPES until the urea concentration in the samples is diluted to two molar.Add trypsin to each sample at a trypsin to protein ratio of one to 50. Incubate at room temperature for at least three hours. Next, add DTT until its concentration is once again one millimolar and incubate at room temperature for two hours.Add iodoacetamide such that its final concentration is 10 millimolar. Incubate in the dark for 30 minutes at room temperature. After this, quench any unreacted iodoacetamide by adding DTT such that its final concentration is 30 millimolar.Incubate at room temperature for 30 minutes. Check the efficiency of the trypsin digestion as outlined in the text protocol. Next, add TFA to each sample such that the final concentration is 1%Using a pH strip, measure the pH of each sample to verify it is between two and three.Add additional TFA drop wise if needed to reach an unacceptable pH. Centrifuge the samples at 20, 000 times g and room temperature for 10 minutes. Collect the supernatant and load the samples onto prepared spin columns.Centrifuge at 100 times g for three minutes or until the sample has passed completely through the column to bind the samples. Next, add 0.5 milliliters of 0.1%TFA to each column. Centrifuge at 500 times g for 30 seconds.After this, add 125 microliters of a solution containing 60%acetonitrile and 0.1 TFA to each column. Centrifuge at 100 times g for three minutes to elute the peptides off the column. Reconstitute each desalted peptide sample in 50 microliters of 50 millimolar HEPES.Using a pH strip, verify that the pH of each sample is between seven and eight. Next, dissolve the TMT reagents in anhydrous acetonitrile, add them to the samples, and then incubate at room temperature for one hour. After this, use 10 microliter pipette tips embedded with chromatography media to desalt one microgram of each sample and analyze the labeling efficiency as outlined in the text protocol.Examine six to 10 separate peptides to ensure that unlabeled peptides are not detected in the labeled samples. Then, mix approximately two microliters of each sample together. Use 10 microliter pipette tips embedded with chromatography media to desalt this mixture.Analyze the samples by LC-MS/MS as outlined in the text protocol. To begin, solubilize the desalted pooled TMT-labeled peptide sample in 65 microliters of buffer A.Use a pH strip to verify that the pH is approximately 8.0. If the sample is still acidic, use ammonium hydroxide to adjust the pH as necessary.Next, fractionate the sample as outlined in the text protocol. Set the fraction collector to collect fractions every two minutes including loading time. Set the flow rate to 0.4 milliliters per minute.Collect a total of 80 fractions. Then, use a vacuum concentrator to completely dry every other sample. Using LC-MS/MS, analyze all 80 fractions to achieve ultra deep proteome coverage.First, pack 1.9 micrometer C18 resin into 75 micrometer inner diameter empty columns reaching a bed volume of approximately 1.3 microliters. Coil the column two to three times inside a butterfly portfolio heater to ensure the entire length is heated. Tape the column to the inside of the heater making sure not to tape over the heater's temperature sensor.Then, heat the column to 65 degrees Celsius. Run 100 nanograms of rat brain peptides through the LC-MS/MS system to assess the system's quality. Repeat this assessment one additional time.After this, load approximately 0.2 micrograms of reconstituted peptides on the column while flowing 5%buffer A.Elute the peptides off the column and analyze with the mass spectrometer as outlined in the text protocol. In this study, a high throughput method for the quantitation of proteins with a 10-plex isobaric labeling strategy is described. TMT labeling efficiency is examined using LC-MS/MS to analyze TMT-labeled samples and corresponding unlabeled samples.As can be seen, the unlabeled peptide peak is found only in the unlabeled sample while the TMT-labeled peptide peak is found only in the labeled sample. Next, the effect of the MS2 isolation window on interference is evaluated. All MS2 scans are designated either as a clean or a noisy scan.While the clean scans exhibit y1 ions of only lysine, the noisy scans exhibit them in both lysine and arginine. Representative interference removal using E.coli peptides individually labeled with three different TMT reagents is shown here. The summed relative intensities reveal that y1 ion-based correction is more accurate for TMT-based quantitation.Once mastered, this protocol can be completed in four to five weeks if done properly. While attempting this procedure, it is very important to make sure all the quality control steps are done completely to ensure the final dataset is as accurate as possible. Adapting this procedure to focus on post-translational modifications can answer other questions such as how ubiquitination, phosphorylation, or acetylation is changing in response to disease progression or drug treatment.After its development, the technology paved the way for researchers in the product field to explore the protein changes in cells and tissues. After watching this video, you should have a very good understanding of how to identify and accurately quantitate over 10, 000 proteins from a mammalian cell or tissue. Don't forget that working with high-pressure UPLC and few silica columns can be extremely hazardous and precautions such as wearing safety goggles should always be taken while performing this procedure.We first had the idea of this method when we began to address the known problem of ratio compression which occurs in isobaric labeling. We have realized extensive fractionation is a great way to not only alleviate this problem, but also increases the identification of proteins and peptides which can further increase and adapts of a proteome analysis.

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Research

JoVE Journal

Biochemistry

Sheathless Capillary Electrophoresis&#8211;Mass Spectrometry for Metabolic Profiling of Biological Samples

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a protocol is presented for the analysis of anionic and cationic metabolites in biological samples by capillary electrophoresis&#8211;mass spectrometry (CE-MS). CE is well-suited for the analysis of highly polar and charged metabolites as compounds are separated on the basis of their charge-to-size ratio. A recently developed sheathless interfacing design, i.e., a porous tip interface, is used for coupling CE to electrospray ionization (ESI) MS. This interfacing approach allows the effective use of the intrinsically low-flow property of CE in combination with MS, resulting in nanomolar detection limits for a broad range of polar metabolite classes. The protocol presented here is based on employing a bare fused-silica capillary with a porous tip emitter at low-pH separation conditions for the analysis of a broad array of metabolite classes in biological samples. It is demonstrated that the same sheathless CE-MS method can be used for the profiling of cationic metabolites, including amino acids, nucleosides and small peptides, or anionic metabolites, including sugar phosphates, nucleotides and organic acids, by only switching the MS detection and separation voltage polarity. Highly information-rich metabolic profiles in various biological samples, such as urine, cerebrospinal fluid and extracts of the glioblastoma cell line, can be obtained by this protocol in less than 1 hr of CE-MS analysis.

A protocol for metabolic profiling of biological samples by capillary electrophoresis&#8211;mass spectrometry using a sheathless porous tip interface design is presented.

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a ...

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

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.

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 ...

Detection of Protein Ubiquitination Sites by Peptide Enrichment and Mass Spectrometry

The posttranslational modification of proteins by the small protein ubiquitin is involved in many cellular events. After tryptic digestion of ubiquitinated proteins, peptides with a diglycine remnant conjugated to the epsilon amino group of lysine (&#39;K-&#949;-diglycine&#39; or simply &#39;diGly&#39;) can be used to track back the original modification site. Efficient immunopurification of diGly peptides combined with sensitive detection by mass spectrometry has resulted in a huge increase in the number of ubiquitination sites identified up to date. We have made several improvements to this workflow, including offline high pH reverse-phase fractionation of peptides prior to the enrichment procedure, and the inclusion of more advanced peptide fragmentation settings in the ion routing multipole. Also, more efficient cleanup of the sample using a filter-based plug in order to retain the antibody beads results in a greater specificity for diGly peptides. These improvements result in the routine detection of more than 23,000 diGly peptides from human cervical cancer cells (HeLa) cell lysates upon proteasome inhibition in the cell. We show the efficacy of this strategy for in-depth analysis of the ubiquitinome profiles of several different cell types and of in vivo samples, such as brain tissue. This study presents an original addition to the toolbox for protein ubiquitination analysis to uncover the deep cellular ubiquitinome.

We present a method for the purification, detection, and identification of diGly peptides that originate from ubiquitinated proteins from complex biological samples. The presented method is reproducible, robust, and outperforms published methods with respect to the level of depth of the ubiquitinome analysis.

The posttranslational modification of proteins by the small protein ubiquitin is involved in many cellular events. After tryptic digestion of ...

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 ...

Nitropeptide Profiling and Identification Illustrated by Angiotensin II

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of reactive oxygen species (ROS) and reactive nitrogen species (RNS) in eukaryotic cells. Precise identification of nitration sites on proteins is crucial for understanding the physiological and pathological processes related to protein nitration, such as inflammation, aging, and cancer. Since the nitrated proteins are of low abundance in cells even under induced conditions, no universal and efficient methods have been developed for the profiling and identification of protein nitration sites. Here we describe a protocol for nitropeptide enrichment by using a chemical reduction reaction and biotin labeling, followed by high resolution mass spectrometry. In our method, nitropeptide derivatives can be identified with high accuracy. Our method exhibits two advantages compared to the previously reported methods. First, dimethyl labeling is used to block the primary amine on nitropeptides, which can be used to generate quantitative results. Second, a disulfide bond containing NHS-biotin reagent is used for the enrichment, which can be further reduced and alkylated to enhance the detection signal on a mass spectrometer. This protocol has been successfully applied to the model peptide Angiotensin II in the current paper.

Proteomic profiling of tyrosine-nitrated proteins has been a challenging technique due to the low abundance of the 3-nitrotyrosine modification. Here we describe a novel approach for nitropeptide enrichment and profiling by using Angiotensin II as the model. This method can be extended for other in vitro or in vivo systems.

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of ...

Absolute Quantification of Cell-Free Protein Synthesis Metabolism by Reversed-Phase Liquid Chromatography-Mass Spectrometry

Cell-free protein synthesis (CFPS) is an emerging technology in systems and synthetic biology for the in vitro production of proteins. However, if CFPS is going to move beyond the laboratory and become a widespread and standard just in time manufacturing technology, we must understand the performance limits of these systems. Toward this question, we developed a robust protocol to quantify 40 compounds involved in glycolysis, the pentose phosphate pathway, the tricarboxylic acid cycle, energy metabolism and cofactor regeneration in CFPS reactions. The method uses internal standards tagged with 13C-aniline, while compounds in the sample are derivatized with 12C-aniline. The internal standards and sample were mixed and analyzed by reversed-phase liquid chromatography-mass spectrometry (LC/MS). The co-elution of compounds eliminated ion suppression, allowing the accurate quantification of metabolite concentrations over 2-3 orders of magnitude where the average correlation coefficient was 0.988. Five of the forty compounds were untagged with aniline, however, they were still detected in the CFPS sample and quantified with a standard curve method. The chromatographic run takes approximately 10 min to complete. Taken together, we developed a fast, robust method to separate and accurately quantify 40 compounds involved in CFPS in a single LC/MS run. The method is a comprehensive and accurate approach to characterize cell-free metabolism, so that ultimately, we can understand and improve the yield, productivity and energy efficiency of cell-free systems.

Here, we present a robust protocol to quantify 40 compounds involved in central carbon and energy metabolism in cell-free protein synthesis reactions. The cell-free synthesis mixture is derivatized with aniline for effective separation using reversed-phase liquid chromatography and then quantified by mass spectrometry using isotopically labelled internal standards.

Cell-free protein synthesis (CFPS) is an emerging technology in systems and synthetic biology for the in vitro production of proteins. However, if CFPS is ...

Label-Free Immunoprecipitation Mass Spectrometry Workflow for Large-scale Nuclear Interactome Profiling

Immunoaffinity purification mass spectrometry (IP-MS) has emerged as a robust quantitative method of identifying protein-protein interactions. This publication presents a complete interaction proteomics workflow designed for identifying low abundance protein-protein interactions from the nucleus that could also be applied to other subcellular compartments. This workflow includes subcellular fractionation, immunoprecipitation, sample preparation, offline cleanup, single-shot label-free mass spectrometry, and downstream computational analysis and data visualization. Our protocol is optimized for detecting compartmentalized, low abundance interactions that are difficult to identify from whole cell lysates (e.g., transcription factor interactions in the nucleus) by immunoprecipitation of endogenous proteins from fractionated subcellular compartments. The sample preparation pipeline outlined here provides detailed instructions for the preparation of HeLa cell nuclear extract, immunoaffinity purification of endogenous bait protein, and quantitative mass spectrometry analysis. We also discuss methodological considerations for performing large-scale immunoprecipitation in mass spectrometry-based interaction profiling experiments and provide guidelines for evaluating data quality to distinguish true positive protein interactions from nonspecific interactions. This approach is demonstrated here by investigating the nuclear interactome of the CMGC kinase, DYRK1A, a low abundance protein kinase with poorly defined interactions within the nucleus.

Described is a proteomics workflow for identifying protein interaction partners from a nuclear subcellular fraction using immunoaffinity enrichment of a given protein of interest and label-free mass spectrometry. The workflow includes subcellular fractionation, immunoprecipitation, filter aided sample preparation, offline cleanup, mass spectrometry, and a downstream bioinformatics pipeline.

Immunoaffinity purification mass spectrometry (IP-MS) has emerged as a robust quantitative method of identifying protein-protein interactions. This ...

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 ...