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 border="0" cellpadding="0" cellspacing="0" style="width:637px;" width="638">
	<tbody>
		<tr height="24">
			<td height="24" style="height:24px;width:216px;">22x22x0.15 mm coverslip</td>
			<td style="width:119px;">VWR</td>
			<td style="width:136px;">470019-004</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">488 nm Argon laser</td>
			<td style="width:119px;">Coherent</td>
			<td style="width:136px;">Innova 70C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">488 nm dichroic mirror</td>
			<td style="width:119px;">Semrock</td>
			<td style="width:136px;">FF495-Di03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">488 nm emission filter</td>
			<td style="width:119px;">Semrock</td>
			<td style="width:136px;">FF02-520/28</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">560 nm dichroic filter</td>
			<td style="width:119px;">Semrock</td>
			<td style="width:136px;">Di02-R561</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">560 nm emission filter</td>
			<td style="width:119px;">Semrock</td>
			<td style="width:136px;">FF02-617/73</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">560 nm fiber laser</td>
			<td style="width:119px;">MBP Communications</td>
			<td style="width:136px;">F-04306-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">60x oil immersion lens</td>
			<td style="width:119px;">Olympus</td>
			<td style="width:136px;">PLAPON 60x</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Avidin</td>
			<td style="width:119px;">Nacalai-tesque</td>
			<td style="width:136px;">03553-64</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Biotin-PEG-amine</td>
			<td style="width:119px;">Thermo Scientific</td>
			<td style="width:136px;">21346</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Biotinylated Alexa Fluor 488</td>
			<td style="width:119px;">Nanocs</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Borate</td>
			<td style="width:119px;">Nacalai-tesque</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">BSA</td>
			<td style="width:119px;">Sigma-Aldrich</td>
			<td style="width:136px;">A9547</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CHAPS</td>
			<td style="width:119px;">Dojindo</td>
			<td style="width:136px;">C008-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cy3 NHS-ester dye</td>
			<td style="width:119px;">GE Healthcare</td>
			<td style="width:136px;">PA13101</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Dialyzer&#160; - D-tube, 6-8 kDa</td>
			<td style="width:119px;">Merck Millipore</td>
			<td style="width:136px;">71507-M</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DMEM</td>
			<td style="width:119px;">Sigma-Aldrich</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DTT</td>
			<td style="width:119px;">Nacalai-tesque</td>
			<td style="width:136px;">14112-94</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">EDC</td>
			<td style="width:119px;">Nacalai-tesque</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">EMCCD camera</td>
			<td style="width:119px;">Andor</td>
			<td style="width:136px;">IiXon 897</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Epi fluorescence microscope</td>
			<td style="width:119px;">Olympus</td>
			<td style="width:136px;">IX81</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:216px;">Gel viewer</td>
			<td style="width:119px;">GE Healthcare</td>
			<td style="width:136px;">ImageQuant LAS 4000</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Penicillin, streptomycine and Amphoterecin mix</td>
			<td style="width:119px;">Gibco</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Plasma cleaner</td>
			<td style="width:119px;">Diener Electronic</td>
			<td style="width:136px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">SDS</td>
			<td style="width:119px;">Wako</td>
			<td style="width:136px;">NC0792960</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Size purification column 10K</td>
			<td style="width:119px;">Merck Millipore</td>
			<td style="width:136px;">UFC5010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tween 20</td>
			<td style="width:119px;">Sigma-Aldrich</td>
			<td style="width:136px;">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...

Watch this Scientific Journal Video about Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level at JoVE.com

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 ...

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Biochemistry

6.1K Views.  RIKEN. Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

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

Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. Thereby, multiple smFRET measurements are used to localize an unknown dye position inside a protein complex by means of trilateration. In order to obtain quantitative information, the Nano-Positioning System (NPS) uses probabilistic data analysis to combine structural information from X-ray crystallography with single-molecule fluorescence data to calculate not only the most probable position but the complete three-dimensional probability distribution, termed posterior, which indicates the experimental uncertainty. The concept was generalized for the analysis of smFRET networks containing numerous dye molecules. The latest version of NPS, Fast-NPS, features a new algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering that allows for the analysis of large smFRET networks in a comparably short time. Moreover, Fast-NPS allows the calculation of the posterior by choosing one of five different models for each dye, that account for the different spatial and orientational behavior exhibited by the dye molecules due to their local environment.
Here we present a detailed protocol for obtaining smFRET data and applying the Fast-NPS. We provide detailed instructions for the acquisition of the three input parameters of Fast-NPS: the smFRET values, as well as the quantum yield and anisotropy of the dye molecules. Recently, the NPS has been used to elucidate the architecture of an archaeal open promotor complex. This data is used to demonstrate the influence of the five different dye models on the posterior distribution.

We present the setup and experimental procedure to obtain smFRET data from large donor-acceptor networks with a TIRF microscope. The step-by-step analysis of these measurements with the Bayesian inference software Fast-NPS yields high-resolved structural information via the application of adapted dye models.

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. ...

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 ...

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Characterization at the Molecular Level using Robust Biochemical Approaches of a New Kinase Protein

Extensive whole genome sequencing has identified many Open Reading Frames (ORFs) providing many potential proteins. These proteins may have important roles for the cell and may unravel new cellular processes. Among proteins, kinases are major actors as they belong to cell signaling pathways and have the ability to switch on or off many processes crucial to the fate of the cell, such as cell growth, division, differentiation, motility, and death.
In this study, we focused on a new potential kinase protein, LIMK2-1. We demonstrated its existence by Western Blot using a specific antibody. We evaluated its interaction with an upstream regulating protein using coimmunoprecipitation experiments. Coimmunoprecipitation is a very powerful technique able to detect the interaction between two target proteins. It may also be used to detect new partners of a bait protein. The bait protein may be purified either via a tag engineered to its sequence or via an antibody specifically targeting it. These protein complexes may then be separated by SDS-PAGE (Sodium Dodecyl Sulfate PolyAcrylamide Gel) and identified using mass spectrometry. Immunoprecipitated LIMK2-1 was also used to test its kinase activity in vitro by &#947;[32P] ATP labeling. This well-established assay may use many different substrates, and mutated versions of the bait may be used to assess the role of specific residues. The effects of pharmacological agents may also be evaluated since this technique is both highly sensitive and quantitative. Nonetheless, radioactivity handling requires particular caution. Kinase activity may also be assessed with specific antibodies targeting the phospho group of the modified amino acid. These kinds of antibodies are not commercially available for all the phospho modified residues.

We characterized a new kinase protein using robust biochemical approaches: Western Blot analysis with a dedicated specific antibody on different cell lines and tissues, interactions by coimmunoprecipitation experiments, kinase activity detected by Western Blot using a phospho-specific antibody and by &#947;[32P] ATP labeling.

Extensive whole genome sequencing has identified many Open Reading Frames (ORFs) providing many potential proteins. These proteins may have important ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for hundreds of individual restriction endonuclease (REase) molecules in one experiment. This assay does not require protein labeling and can be carried out with a single imaging channel. In addition, the results of multiple individual experiments can be pooled to generate well-populated dwell-time distributions. Analysis of the resulting dwell-time distributions can help elucidate the DNA cleavage mechanism by revealing the presence of kinetic steps that cannot be directly observed. Example data collected using this assay with the well-studied REase, EcoRV - a dimeric Type IIP restriction endonuclease that cleaves the palindromic sequence GAT&#8595;ATC (where &#8595; is the cut site) - are in agreement with prior studies. These results suggest that there are at least three steps in the pathway to DNA cleavage that is initiated by introducing magnesium after EcoRV binds DNA in its absence, with an average rate of 0.17 s-1 for each step.

Using quantum-dot-labeled DNA and total internal reflection fluorescence microscopy, we can investigate the reaction mechanism of restriction endonucleases while using unlabeled protein. This single-molecule technique allows for massively multiplexed observation of individual protein-DNA interactions, and data can be pooled to generate well-populated dwell-time distributions.

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for ...

Making Precise and Accurate Single-Molecule FRET Measurements using the Open-Source smfBox

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes measurements on freely diffusing biomolecules more accessible. This overview includes a step-by-step protocol for using this instrument to make measurements of precise FRET efficiencies in duplex DNA samples, including details of the sample preparation, instrument setup and alignment, data acquisition, and complete analysis routines. The presented approach, which includes how to determine all the correction factors required for accurate FRET-derived distance measurements, builds on a large body of recent collaborative work across the FRET Community, which aims to establish standard protocols and analysis approaches. This protocol, which is easily adaptable to a range of biomolecular systems, adds to the growing efforts in democratising smFRET for the wider scientific community.

This article provides step-by-step instructions for making fully-corrected accurate FRET measurements on individual, freely diffusing biomolecules using the open-source, inexpensive smfBox, from switch on, through alignment and focusing, to data collection and analysis.

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes ...