The authors thank Masae Ohno and Kazuya Nishimura for experimental assistance and advice. This work was supported by PRESTO (JPMJPR15F7), Japan Science and Technology Agency, Grants-in-aid for Young Scientists (A) (24687022), Challenging Exploratory Research (26650055), and Scientific Research on Innovative Areas (23115005), Japan Society for the Promotion of Science, and by grants from the Takeda Science Foundation and the Mochida Memorial Foundation for Medical and Pharmaceutical Research. S.L. acknowledges support from the RIKEN International Program Associate (IPA) program.

1. Cell preparation
<ol>
	<li>Cultivate HeLa cells in a 10 cm dish at 37 &#176;C under 5% CO2 in Dulbecco&#8217;s modified Eagle&#8217;s medium containing 10% fetal bovine serum.</li>
	<li>Collect exponentially growing cells once they have reached 70% confluency, following ATCC instructions17.
	<ol>
		<li>Rinse cells with 5 mL of 1x phosphate buffered saline (PBS) (pH 7.4). Remove PBS.</li>
		<li>Add 1 mL of 0.1% trypsin-ethylenediaminetetraacetic acid (EDTA). Incubate 5 min at 37 &#176;C.</li>
		<li>Monitor the progression of cell dissociation by microscopy.</li>
		<li>Once cells become round and are detached from the dish, add 4 mL of growth medium, and break cell agglomeration by pipetting.</li>
		<li>Count the number of cells using a cell counter.</li>
		<li>Centrifuge at 150 x g for 10 min in a centrifuge tube. Remove the medium and resuspend the cell pellet with 5 mL of 1x PBS (pH 7.4).</li>
		<li>Repeat step 1.2.6. Resuspend cells in 1x PBS (pH 7.4) to a final concentration of 106 cells/mL using the count number in step 1.2.5. 
		NOTE: The cell suspension can be aliquoted and stored at -80 &#176;C for several months. Repeating a freeze/thaw cycle should be avoided.</li>
	</ol>
	</li>
</ol>
2. Cell lysis and fluorescent labeling
<ol>
	<li>Make 10 mL of lysing buffer (0.1 M borate, 1% sodium dodecyl sulfate (SDS) and 1% Tween 20). Adjust to pH 12 using 2 M NaOH. 
	CAUTION: Concentrated NaOH is corrosive. Wear suitable gloves and glasses when preparing this solution. 
	NOTE: Lysing buffer can be stored for up to one week at room temperature.</li>
	<li>Mix 100 &#956;L of lysing buffer with 1 &#956;L of 1 M dithiothreitol (DTT) and 4 &#956;L of 50% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS).</li>
	<li>Mix 1 &#956;L of the cell culture (approximately 1,000 cells) with the mixed lysing buffer from step 2.2. Homogenize the solution by slowly pipetting to avoid bubbles. Incubate in the dark at room temperature for 5 min with gentle agitation. Re-homogenize the solution by pipetting slowly.</li>
	<li>Add 1 &#956;L of 2 &#956;g/mL Cy3 NHS-ester dye. Homogenize the solution by slowly pipetting to avoid bubbles. Incubate for 5 min in the dark at room temperature with gentle agitation. 
	NOTE: The high pH of this buffer causes neutralization of lysine residues in proteins, leading to a higher reactivity.</li>
	<li>Repeat the addition of 1 &#956;L of 2 &#956;g/mL Cy3 NHS-ester dye. Homogenize the solution by slowly pipetting to avoid bubbles. Incubate for 10 min in the dark at room temperature with gentle agitation. 
	NOTE: This repeated addition of NHS-ester dye can increase the labeling efficiency because some of the Cy3 NHS-ester dye is hydrolyzed and deactivated by the high pH of the reaction buffer.</li>
	<li>Adjust to pH 7.2 by adding 100 &#956;L of 0.8 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-NaOH pH 7.2 to quench the reaction. Homogenize the solution by slowly pipetting to avoid bubbles.</li>
	<li>Add 20 &#956;L of 19 mg/mL biotin-(polyethylene glycol (PEG))2-amine and 5 &#956;L of 20 mg/mL 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC). Homogenize the solution by slowly pipetting to avoid bubbles. Incubate for 1 h in the dark at room temperature with gentle agitation. 
	NOTE: This biotinylation step is necessary to fix the protein molecules on a microscope coverslip at a fixed density and with no bias for protein species.</li>
	<li>Remove unreacted labeling reagents and concentrate the sample using an ultrafiltration column with 10 kDa cut-off, following the manufacturer&#39;s instructions. 
	NOTE: The protocol can be paused here. The sample can be stored at -80 &#176;C for several weeks.</li>
</ol>
3. Proteome separation and recovery
<ol>
	<li>Add 4x SDS sample buffer (200 mM Tris-HCl pH 6.8, 4% SDS, 4% glycerol and 0.4% bromophenol blue). 
	NOTE: To avoid dye damage, the protein sample should not be heated as done in standard SDS-PAGE, and should be kept at room temperature.</li>
	<li>Perform standard SDS-PAGE for the protein sample and a molecular ladder, using 20% acrylamide gel. Migrate the sample in the gel at a constant voltage (160 V) until the migration front just exits the gel.</li>
	<li>Image the gel to identify the location of the protein bands (Figure 3).</li>
	<li>Cut out the gel portions including protein fractions of interest using a sharp blade, based on the band locations. Fractioning at 16, 23, 55 and 120 kDa peaks, and at 19&#8211;25, 25&#8211;32, 32&#8211;42, 42&#8211;54, 54&#8211;69, 69&#8211;97, 97&#8211;136 and 136&#8211;226 kDa regions are shown in Figure 3.</li>
	<li>Extract proteins using a dialyzer by electroelution with a pore size of 6&#8211;8 kDa, following the manufacturer&#39;s instructions. 
	NOTE: The extracted sample can be stored at -80 &#176;C for several weeks.</li>
</ol>
4. Microscope coverslip preparation
<ol>
	<li>Prepare 200 &#956;L of avidin buffer (5 mM HEPES-NaOH pH 7.2, 10 ng/mL avidin and 2 mg/mL bovine serum albumin (BSA)) for each sample.</li>
	<li>Prepare coverslips of 22 mm x 22 mm and 0.15 mm thickness. Expose the coverslips to air plasma using a plasma cleaner for 1 min to clean and activate their surfaces.</li>
	<li>Coat the coverslips with avidin by spin-coating 200 &#956;L of avidin buffer using a spin coater for 5 seconds at 500 rpm, followed by 30 seconds at 1,000 rpm. Incubate the coverslips for 15 min at room temperature to let them dry.</li>
	<li>Prepare coverslips for the protein sample.
	<ol>
		<li>Dilute the protein sample (from 3.5) 100 times in 5 mM HEPES-NaOH pH 7.2.</li>
		<li>Place 100 &#956;L of the diluted sample on the surface at the center of the avidin-coated coverslip (from step 4.3).</li>
		<li>Incubate in the dark for 15 min at room temperature.</li>
	</ol>
	</li>
	<li>Prepare the positive control.
	<ol>
		<li>Place 100 &#956;L of 10 ng/mL purified fluorescent biotin in 5 mM HEPES-NaOH pH 7.2 at the center of the avidin coated coverslip (from step 4.3).</li>
		<li>Incubate in the dark for 15 min at room temperature. 
		NOTE: This positive control coverslip provides the density of avidin binding spots, which is used to calculate the LO.</li>
	</ol>
	</li>
	<li>Prepare the negative control.
	<ol>
		<li>Spin-coat the plasma cleaned coverslip with 200 &#956;L of 5 mM HEPES-NaOH and 2 mg/mL BSA (without avidin).</li>
		<li>Incubate for 15 min at room temperature to dry.</li>
		<li>Add 100 &#956;L of 10 ng/mL purified fluorescent biotin in 5 mM HEPES-NaOH pH 7.2.</li>
		<li>Incubate in the dark for 15 min at room temperature. 
		NOTE: This negative control coverslip provides the amount of non-specific binding of fluorescent biotin to the coverslip without avidin.</li>
	</ol>
	</li>
	<li>Rinse each coverslip with 200 &#956;L of distilled water by pipetting at the edges of the coverslip so as not to touch the middle of the coverslip with the tip. Repeat this wash three times. 
	NOTE: After aspirating water on the coverslip, water should be immediately placed back on the coverslip to prevent it from drying out completely.</li>
	<li>Expose air plasma to the same number of coverslips as in step 4.2.</li>
	<li>Place the cleaned coverslip on the sample-bound coverslip from step 4.7 to avoid drying.</li>
</ol>
5. Observation with single molecule fluorescence microscopy
<ol>
	<li>Start up a wide-field or evanescent field fluorescence microscope. 
	CAUTION: Direct or scattered laser radiation can cause eye or skin damage. Wear suitable protective goggles and shield the laser light path. 
	NOTE: For the microscope setup, please refer to our previous publication16. Briefly, a wide-field or evanescent field fluorescence microscope equipped with a high-power 488 nm laser excitation, a high numerical aperture objective lens and a high sensitivity camera can be used for this measurement. Further, for an automated measurement, a computer-controlled XY translational sample stage, mechanical shutters for laser excitations and an auto-focusing system are recommended to be installed.</li>
	<li>Set the coverslip on the microscope and find the focus.</li>
	<li>Perform a tile scan to obtain at least 100 images. 
	NOTE: The laser illumination should be controlled with mechanical shutters to be exposed to the sample only when recording images with the camera, to minimize photobleaching. If there are many samples, automated measurement using a computer program to control the microscope devices is recommended. Each image typically includes 100&#8211;500 single molecule spots when using a 60x objective lens.</li>
</ol>
6. Image analysis and extraction of information
<ol>
	<li>If the image has unevenness due to the laser illumination pattern, carry out image processing to correct it18.
	<ol>
		<li>Acquire a reference image by measuring with a coverslip sample consisting of uniformly-spread dyes using a camera setting the same as in step 5.3.</li>
		<li>Acquire an offset image by measuring with no laser excitation using the same camera setting.</li>
		<li>Subtract each pixel value in every sample image and the reference image with the value of the offset image.</li>
		<li>Divide each pixel value in the subtracted sample images by the value in the subtracted reference image.</li>
	</ol>
	</li>
	<li>Collect properly acquired images and remove the image background.
	<ol>
		<li>Remove images containing large fluorescent aggregates manually.</li>
		<li>Remove the image background by applying the rolling-ball algorithm19 with a ball radius of 50 pixels using ImageJ20.</li>
		<li>Sharpen images by subtracting blurred images using ImageJ (sharpen image function).</li>
	</ol>
	</li>
	<li>Find and analyze single molecule spots in images.
	<ol>
		<li>Identify single molecule spots with the Yen threshold algorithm21 using ImageJ.</li>
		<li>Filter spots with less than two pixels to exclude dark noise of the camera and spots more than 20 pixels to exclude aggregations of proteins using ImageJ (ROI manager).</li>
		<li>Count the number of spots in each image. Using the illumination corrected images (step 6.1.4), analyze total intensities within pixels for every spot using ImageJ.</li>
	</ol>
	</li>
	<li>Calculate the labeling occupancy (LO) using the following formula: 
	LO = (number of counts in the sample / number of counts in the positive control) x 100</li>
	<li>Calculate the average number of fluorescent dyes bound to a single protein molecule (n&#772;dye).
	<ol>
		<li>Analyze a histogram of intensities of every spot. 
		NOTE: The histogram usually has several peaks, and every peak represents a different number of dye molecules bound to a protein. The first, second and third peak in the histogram corresponds to 1, 2 and 3 molecules, respectively.</li>
		<li>Calculate the average intensity from the histogram. Also calculate the peak intensity of the first peak by applying a Gaussian fitting.</li>
		<li>Calculate n&#772;dye by dividing the average intensity with the peak intensity.</li>
	</ol>
	</li>
</ol>

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The authors declare the following competing financial interest(s): RIKEN has filed a patent application on these results with S.L. and Y.T. named as co-inventors.

This paper describes a protocol to quantify the labeling homogeneity of each labeled protein species in cells after separation with SDS-PAGE (step 3). The separation method can be substituted with other methods such as liquid chromatography or capillary electrophoresis, which allow separation and fractionation of cellular proteins with high resolution, while requiring special equipment23. The labeling method using NHS-ester in the current protocol can be replaced with other methods using maleimide...

Proteome analysis, which aims to quantify the entire set of protein molecules expressed in the cell, is a valuable approach in current biological and medicinal studies. This analysis commonly relies on mass spectrometry, which identifies protein species based on spectra generated through protein ionization1,2,3. An alternative method for proteome analysis is electrophoresis, including polyacrylamide gel electrophoresis (PAGE), capillary electrophoresis and two-dimensional (2D) gel electrophoresis. This method relies on non-specific fluorescent labeling of all the protein molecules in the analyzed cells, followed by electrophoretic separation and detection and quantification of each protein species. To achieve the required non-specific protein labeling, one strategy is to use fluorescent dyes that can bind to proteins via electrostatic and hydrophobic interactions, such as Coomassie Blue and Sypro-Ruby4,5,6. An alternative strategy is to use covalent labeling with dyes containing N-hydroxysuccinimide (NHS) ester or maleimide, which can covalently bind to proteins through common residues such as primary amines and thiols, respectively7,8.
Meanwhile, the sensitivity of fluorescence detection is ideal for analyzing low-abundance proteins and small numbers of cells. Single molecule fluorescence imaging is one of the most sensitive methods that allows the detection of individual fluorescent dyes and labeled proteins in vitro and in vivo9,10,11,12,13,14,15. Application of this imaging method to electrophoresis-based proteome analysis is expected to enable highly sensitive and quantitative assays by counting individual fluorescently-labeled proteins. However, it remains unclear whether labeling with fluorescent dyes is homogeneous enough across all the protein molecules, and how this homogeneity is affected by different protein species (Figure 1). Simple bulk solution measurements can be used to obtain a molar ratio of fluorescent dyes to proteins called &#8216;coupling efficiency&#8217;8 or &#8216;labeling efficiency&#8217;, but this property does not provide information on the homogeneity of the labeling among protein molecules.
Here, we describe the protocol for an assay to investigate the labeling homogeneity for all protein species in the cell (Figure 2)16. The two key steps of this assay are protein purification and imaging. In the first step, all the proteins in the cells are fluorescently labeled and biotinylated, then extracted separately using gel electrophoresis followed by electroelution. In the second step, fluorescence properties of individual protein molecules in the extracted samples are evaluated based on single molecule imaging. From this data, parameters important for the counting analysis, such as the percentage of proteins labeled with a least one dye, which we call labeling occupancy (LO)16, and the average number of fluorescent dyes bound to a single protein molecule (n&#772;dye), can be characterized. In the protocol, an optimized procedure for labeling the HeLa cell proteome with NHS ester-based Cyanine 3 (Cy3) dye is presented as an example, and can be modified with other labeling procedures according to desired research goals.

<table><tbody><tr><td>22x22x0.15 mm coverslip</td><td>VWR</td><td>470019-004</td><td></td></tr><tr><td>488 nm Argon laser</td><td>Coherent</td><td>Innova 70C</td><td></td></tr><tr><td>488 nm dichroic mirror</td><td>Semrock</td><td>FF495-Di03</td><td></td></tr><tr><td>488 nm emission filter</td><td>Semrock</td><td>FF02-520/28</td><td></td></tr><tr><td>560 nm dichroic filter</td><td>Semrock</td><td>Di02-R561</td><td></td></tr><tr><td>560 nm emission filter</td><td>Semrock</td><td>FF02-617/73</td><td></td></tr><tr><td>560 nm fiber laser</td><td>MBP Communications</td><td>F-04306-2</td><td></td></tr><tr><td>60x oil immersion lens</td><td>Olympus</td><td>PLAPON 60x</td><td></td></tr><tr><td>Avidin</td><td>Nacalai-tesque</td><td>03553-64</td><td></td></tr><tr><td>Biotin-PEG-amine</td><td>Thermo Scientific</td><td>21346</td><td></td></tr><tr><td>Biotinylated Alexa Fluor 488</td><td>Nanocs</td><td></td><td></td></tr><tr><td>Borate</td><td>Nacalai-tesque</td><td></td><td></td></tr><tr><td>BSA</td><td>Sigma-Aldrich</td><td>A9547</td><td></td></tr><tr><td>CHAPS</td><td>Dojindo</td><td>C008-10</td><td></td></tr><tr><td>Cy3 NHS-ester dye</td><td>GE Healthcare</td><td>PA13101</td><td></td></tr><tr><td>Dialyzer&amp;nbsp; - D-tube, 6-8 kDa</td><td>Merck Millipore</td><td>71507-M</td><td></td></tr><tr><td>DMEM</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>DTT</td><td>Nacalai-tesque</td><td>14112-94</td><td></td></tr><tr><td>EDC</td><td>Nacalai-tesque</td><td></td><td></td></tr><tr><td>EMCCD camera</td><td>Andor</td><td>IiXon 897</td><td></td></tr><tr><td>Epi fluorescence microscope</td><td>Olympus</td><td>IX81</td><td></td></tr><tr><td>Gel viewer</td><td>GE Healthcare</td><td>ImageQuant LAS 4000</td><td></td></tr><tr><td>Penicillin, streptomycine and Amphoterecin mix</td><td>Gibco</td><td></td><td></td></tr><tr><td>Plasma cleaner</td><td>Diener Electronic</td><td></td><td></td></tr><tr><td>SDS</td><td>Wako</td><td>NC0792960</td><td></td></tr><tr><td>Size purification column 10K</td><td>Merck Millipore</td><td>UFC5010</td><td></td></tr><tr><td>Tween 20</td><td>Sigma-Aldrich</td><td>P9416</td><td></td></tr></tbody></table>

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.

Figure 4 represents raw image data for different molecular weight fractions of proteins from HeLa cell lysate, as well as the positive and negative control. While both the protein sample and positive control exhibit 100&#8211;500 spots per image, the negative control displays none or a few spots, showing that the protocol sufficiently inhibits non-specific binding of dyes to the coverslip surface. Spot intensity histograms for the protein samples and the posi...

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

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 ...

proteome-wide-quantification-labeling-homogeneity-at-single-molecule

Research

JoVE Journal

Biochemistry

6.1K Views.  RIKEN. Deze methode toont de kalibratie van het kwantificeren van eiwithoeveelheid in vloeibaar monster met single-molecule proces voor het tellen. Specifiek, het kan illustreren hoe eiwit populaties fluorescerend kunnen worden gelabeld op elk molecuul niveau. Eerdere methoden uitgevoerd bulk metingen om molaire verhoudingen van fluorescerende en eiwitmoleculen te analyseren, maar ze kunnen niet onthullen hoe heterogeen eiwit populaties daadwerkelijk worden gelabeld.Onze methoden kunnen het d...

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

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) ...

A Quantitative Glycomics and Proteomics Combined Purification Strategy

There is a growing desire in the biological and clinical sciences to integrate and correlate multiple classes of biomolecules to unravel biology, define pathways, improve treatment, understand disease, and aid biomarker discovery. N-linked glycosylation is one of the most important and robust post-translational modifications on proteins and regulates critical cell functions such as signaling, adhesion, and enzymatic function. Analytical techniques to purify and analyze N-glycans have remained relatively static over the last decade. While accurate and effective, they commonly require significant expertise and resources. Though some high-throughput purification schemes have been developed, they have yet to find widespread adoption and often rely on the enrichment of glycopeptides. One promising method, developed by Thomas-Oates et al., filter aided N-glycan separation (FANGS), was qualitatively demonstrated on tissues. Herein, we adapted FANGS to plasma and coupled it to the individuality normalization when labeling with glycan hydrazide tags strategy in order to achieve accurate relative quantification by liquid chromatography mass spectrometry and enhanced electrospray ionization. Furthermore, we designed new functionality to the protocol by achieving tandem, shotgun proteomics and glycosylation site analysis on hen plasma. We showed that N-glycans purified on filter and derivatized by hydrophobic hydrazide tags were comparable in terms of abundance and class to those by solid phase extraction (SPE); the latter is considered a gold standard in the field. Importantly, the variability in the two protocols was not statistically different. Proteomic data that was collected in-line with glycomic data had the same depth compared to a standard trypsin digest. Peptide deamidation is minimized in the protocol, limiting non-specific deamidation detected at glycosylation motifs. This allowed for direct glycosylation site analysis, though the protocol can accommodate 18O site labeling as well. Overall, we demonstrated a new in-line high-throughput, unbiased, filter based protocol for quantitative glycomics and proteomics analysis.

A high-throughput protocol was developed for combined proteomics and glycomics purification and LC-MS/MS quantification in plasma. Deamidation analysis of N-linked glycosylation motifs was specific to deglycosylated sites. Accurate quantitation of N-glycans was achieved by coupling filter aided N-glycan separation to the individuality normalization when labeling with glycan hydrazide tags strategy.

There is a growing desire in the biological and clinical sciences to integrate and correlate multiple classes of biomolecules to unravel biology, define ...

Quantitative Analysis of Chromatin Proteomes in Disease

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due to the high percentage of binucleated cells,1 the predominantly heterochromatic state of the DNA, and the non-dividing nature of the cardiomyocyte which renders adult nuclei in a permanent state of interphase.2 Transcriptional regulation during development and disease have been well studied in this organ,3-5 but what remains relatively unexplored is the role played by the nuclear proteins responsible for DNA packaging and expression, and how these proteins control changes in transcriptional programs that occur during disease.6 In the developed world, heart disease is the number one cause of mortality for both men and women.7 Insight on how nuclear proteins cooperate to regulate the progression of this disease is critical for advancing the current treatment options.
Mass spectrometry is the ideal tool for addressing these questions as it allows for an unbiased annotation of the nuclear proteome and relative quantification for how the abundance of these proteins changes with disease. While there have been several proteomic studies for mammalian nuclear protein complexes,8-13 until recently14 there has been only one study examining the cardiac nuclear proteome, and it considered the entire nucleus, rather than exploring the proteome at the level of nuclear sub compartments.15 In large part, this shortage of work is due to the difficulty of isolating cardiac nuclei. Cardiac nuclei occur within a rigid and dense actin-myosin apparatus to which they are connected via multiple extensions from the endoplasmic reticulum, to the extent that myocyte contraction alters their overall shape.16 Additionally, cardiomyocytes are 40% mitochondria by volume17 which necessitates enrichment of the nucleus apart from the other organelles. Here we describe a protocol for cardiac nuclear enrichment and further fractionation into biologically-relevant compartments. Furthermore, we detail methods for label-free quantitative mass spectrometric dissection of these fractions-techniques amenable to in vivo experimentation in various animal models and organ systems where metabolic labeling is not feasible.

Advances in mass spectrometry have allowed the high throughput analysis of protein expression and modification in a host of tissues. Combined with subcellular fractionation and disease models, quantitative mass spectrometry and bioinformatics can reveal new properties in biological systems. The method described herein analyzes chromatin-associated proteins in the setting of heart disease and is readily applicable to other in vivo models of human disease.

In the nucleus reside the proteomes whose functions are most intimately linked with gene regulation. Adult mammalian cardiomyocyte nuclei are unique due ...

Medicine

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

"Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome

Over the past decade, mass spectrometry-based proteomics has enabled an in-depth characterization of biological systems across a broad array of applications. The cell surface proteome ("surfaceome") in human disease is of significant interest, as plasma membrane proteins are the primary target of most clinically approved therapeutics, as well as a key feature by which to diagnostically distinguish diseased cells from healthy tissues. However, focused characterization of membrane and surface proteins of the cell has remained challenging, primarily due to the complexity of cellular lysates, which mask proteins of interest by other high-abundance proteins. To overcome this technical barrier and accurately define the cell surface proteome of various cell types using mass spectrometry proteomics, it is necessary to enrich the cell lysate for cell surface proteins prior to analysis on the mass spectrometer. This paper presents a detailed workflow for labeling cell surface proteins from cancer cells, enriching these proteins out of the cell lysate, and subsequent sample preparation for mass spectrometry analysis.

Here, we describe a proteomics workflow for characterization of the cell surface proteome of various cell types. This workflow includes cell surface protein enrichment, subsequent sample preparation, analysis using an LC-MS/MS platform, and data processing with specialized software.

Over the past decade, mass spectrometry-based proteomics has enabled an in-depth characterization of biological systems across a broad array of ...

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex composition under different conditions. In this protocol the approach is demonstrated by comparing the composition of the protein complex responsible for cyclic electron flow (CEF) in Chlamydomonas reinhardtii, isolated from genetically different strains. The procedure comprises the isolation of thylakoid membranes, followed by their separation into multiprotein complexes by sucrose density gradient centrifugation, SDS-PAGE, immunodetection and comparative, quantitative mass spectrometry (MS) based on differential metabolic labeling (14N/15N) of the analyzed strains. Detergent solubilized thylakoid membranes are loaded on sucrose density gradients at equal chlorophyll concentration. After ultracentrifugation, the gradients are separated into fractions, which are analyzed by mass-spectrometry based on equal volume. This approach allows the investigation of the composition within the gradient fractions and moreover to analyze the migration behavior of different proteins, especially focusing on ANR1, CAS, and PGRL1. Furthermore, this method is demonstrated by confirming the results with immunoblotting and additionally by supporting the findings from previous studies (the identification and PSI-dependent migration of proteins that were previously described to be part of the CEF-supercomplex such as PGRL1, FNR, and cyt f). Notably, this approach is applicable to address a broad range of questions for which this protocol can be adopted and e.g. used for comparative analyses of multiprotein complex composition isolated from distinct environmental conditions.

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The described comparative, quantitative proteomic approach aims at obtaining insights into the composition of multiprotein complexes&#160;under different conditions and is demonstrated by comparing genetically different strains. For quantitative analysis equal volumes of different fractions from a sucrose density gradient are mixed and analyzed by mass spectrometry.

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex ...

Biology

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ ...

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 ...

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

Samenvatting

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