The authors thank Dr. Wolf-Dieter Stalz, Dr. Arnaud Remy, Dr. Anton Posch, Dr. Patricia Piatti, Tom Davies, Kris Simonyi, and Jeff Durban for their critical review and editing of this manuscript. The authors also thank Allison Schwartz for technical support.

1. Protein Sample Prep
(A typical procedure to extract proteins from cell culture is described)
<ol>
	<li>Place the HeLa cell culture dish in ice and wash the cells with ice-cold Tris-buffered Saline (TBS; 20 mM Tris-HCl, pH 7.5, 150 mM NaCl).</li>
	<li>Aspirate the TBS, then add 1 ml per 100 mm dish ice-cold RIPA buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) supplemented with phosphatase and protease inhibitors.</li>
	<li>Scrape adherent cells off the dish using a cold plastic cell scraper; gently transfer the cell suspension into a precooled microcentrifuge tube.</li>
	<li>Maintain constant agitation for 30 min at 4 &#176;C on a rotator.</li>
	<li>Spin at 16,000 x g for 20 min in a 4 &#176;C precooled centrifuge.</li>
	<li>Gently remove the tube from the centrifuge and place on ice. Transfer the supernatant to a fresh tube kept on ice, and discard the pellet.</li>
	<li>Remove a small volume (10-20 &#956;l) of lysate to perform a protein assay. Determine the protein concentration for each sample using the RC DC assay kit.</li>
	<li>If needed, aliquot the protein samples for long term storage at -20 &#176;C. Repeat freeze and thaw cycles cause protein degradation and should be avoided.</li>
	<li>Take about 20 &#956;g of each sample, add an equal volume of 2x Laemmli Sample Buffer (4% SDS, 10% 2-mercaptoethanol, 20% glycerol, 0.004% bromophenol blue, 125 mM Tris-HCl, pH 6.8).</li>
	<li>Heat each cell lysate in sample buffer at 95 &#176;C for 5 min.</li>
	<li>Centrifuge at 16,000 x g in a microcentrifuge for 1 min.</li>
</ol>
2. Gel Electrophoresis with Stain-free Gels (~30 min)
<ol>
	<li>Take a Criterion TGX Any KD stain-free precast gel (a midi format gel), remove the comb and the tape from the bottom of the cassette.</li>
	<li>Place the cassette in the Criterion cell and fill the integrated upper buffer chamber with 60 ml running buffer (25 mM Tris, 190 mM glycine, 0.1% SDS, pH 8.3). Rinse the wells with running buffer.</li>
	<li>Fill each half of the lower buffer tank with 400 ml running buffer to the marked fill line.</li>
	<li>Load the protein samples and appropriate protein markers.</li>
	<li>Place the lid on the tank, aligning the color-coded banana plugs with corresponding jacks on the lid.</li>
	<li>Run the gel for ~30 min at 200 V or 20 min at 300 V.</li>
</ol>
3. Stain-free Gel Imaging Using the Chemidoc MP System to Check Protein Separation Quality (~5 min)
<ol>
	<li>Remove the gel cassette from the cell. Use the gel cassette opening tool in the Criterion Cell lid to crack open the cassette and release the gel.</li>
	<li>Apply a couple of milliliters of water to the center of the UV sample tray of the ChemiDoc MP imager. Carefully lift the gel from the cassette and place it on the tray.</li>
	<li>Launch Image Lab software and capture the stain-free gel image (Figure 1A) with the following settings: 
	Application: stain-free gel 
	Gel activation time: 1 min 
	Imaging area: criterion gel 
	Image exposure time: automatically optimized for the most intense bands</li>
	<li>Remove the gel from the sample tray and proceed immediately to transfer step.</li>
</ol>
4. Protein Transfer with the Trans-Blot Turbo System (~10 min)
<ol>
	<li>Open a Trans-Blot Turbo Midi PVDF Transfer Pack; place the bottom stack (which includes the membrane) on the base of the transfer cassette.</li>
	<li>Place the gel on top of the membrane, place the top stack on the gel, and roll out bubbles.</li>
	<li>Place the lid on the cassette base and turn the dial to lock it.</li>
	<li>Insert the cassette into the blotter bay.</li>
	<li>Start the transfer by selecting a preset Turbo program and choosing the Criterion gel size (midi), and then press RUN. A typical run takes only 7 min.</li>
	<li>When the transfer is over, disassemble the blotting sandwich and place both the blot and the gel in a container with deionized water.</li>
</ol>
5. Stain-free Gel and Blot Imaging Using the Chemidoc MP System to Check Protein Transfer Efficiency and Quality (~5 min)
<ol>
	<li>Place the post-transfer gel on the sample tray of the ChemiDoc MP imager.</li>
	<li>Launch Image Lab software and capture the stain-free image of the post-transfer gel (Figure 1B) with the following settings: 
	Application: stain-free gel 
	Gel activation time: none 
	Imaging area: criterion gel 
	Image exposure time: same to the exposure time for the pretransfer gel image</li>
	<li>Remove the gel from the sample tray and then image the blot (Figure 1C) with the following settings. Keep the blot wet with a few drops of water or TBST when imaging. 
	Application: stain-free blot 
	Imaging area: criterion gel 
	Image exposure time: automatically optimized for the most intense bands</li>
	<li>Remove the blotting membrane from the sample tray and place it in a container with TBST (0.1% Tween 20 in TBS).</li>
</ol>
6. Antibody Incubation
<ol>
	<li>Block by placing the blot in a solution of 3% bovine serum albumin (BSA) in TBST at room temp for 1 hr.</li>
	<li>Incubate the blot overnight at 4 &#176;C in the solution containing mouse primary antibody raised against the first target protein and rabbit primary antibody raised against the second target protein.</li>
	<li>Pour out the solution containing the primary antibody. Next, wash the blot by agitating in 20 ml of TBST for 5 min. Repeat 4x for a total of 5 washes.</li>
	<li>Incubate for 1 hr at room temp in the secondary antibody solution containing a Dylight 650 conjugated Goat-anti-mouse antibody and a Dylight 549 conjugated Goat-anti-rabbit antibody.</li>
	<li>Pour out the solution containing the primary antibody. Next, wash the blot by agitating in 20 ml of TBST for 5 min. Repeat 4x for a total of 5 washes.</li>
</ol>
7. Imaging and Data Analysis by Image Lab Software- Total Protein Normalization (~5 min)
<ol>
	<li>Acquire a multiplexing fluorescent image of the blot (Figure 1E) by opening a new multichannel protocol, configure three fluorescent channels and run the protocol. 
	Channel 1: 
	Application: blot Dylight 650 
	Imaging area: criterion gel 
	Image exposure time: automatically optimized for the most intense bands 
	Channel 2: 
	Application: blot Dylight 549 
	Imaging area: criterion gel 
	Image exposure time: automatically optimized for the most intense bands 
	Channel 3: 
	Application: stain-free blot 
	Imaging area: criterion gel Image exposure time: automatically optimized for the most intense bands</li>
	<li>Click the Normalization icon from the Analysis Tool Box and click Yes to detect lanes and bands.</li>
	<li>Select and use &#34;Lanes and Bands tools&#34; to make adjustments to the lanes and bands if needed.</li>
	<li>Select stain-free image as the normalization channel.</li>
	<li>Select MW Analysis Tools and assign the MW standard lanes by checking the boxes below them.</li>
	<li>To view the normalized volumes, click the Analysis Table on the tool bar. All calculations will be performed automatically by the software, including the Normalization Factor and Normalized Volumes. The target protein band intensity values are now adjusted for variation in the protein load. This will allow accurate comparisons of target proteins among the samples.</li>
</ol>

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The authors, Anton Posch, Jonathan Kohn, Kenneth Oh, Matt Hammond, and Ning Liu are employees of Bio-Rad Laboratories, Inc.

The V3 stain-free protocol described above is for multiplexing fluorescent western blotting. It can also be applied in western blotting using chemiluminescent detection. In the multiplex fluorescent western blot protocol, the stain-free blot images are acquired at two time points: 1) right after protein transfer; 2) at the multiplex fluorescent imaging step after antibody incubation. The first stain free image is used to calculate transfer efficiency and the second stain-free image is used as the loading control. When the chemiluminescent method is applied, it is not possible to take a multiplex image for the stain-free and target protein signals, because the chemiluminescent signal will show up in the stain-free channel too. In this case, we recommend to use the stain-free image taken right after the protein transfer step for loading control and normalization analysis.
The V3 western workflow provides the following unique benefits compared to the traditional western blot workflow that uses housekeeping proteins as loading controls:
First, the V3 workflow provides a practical, convenient and more reliable loading control to validate the changes in the level of the protein of interest. The V3 protocol uses a total protein loading control to normalize the level of the protein of interest measured in each sample. It avoids two pitfalls of using the housekeeping proteins as loading controls: saturated immunodetection and inconsistent housekeeping protein expression levels among the samples under certain experimental conditions. Stripping and reprobing steps with antibodies directed against housekeeping are no longer necessary. Using stain-free technology, there is no need to stain and de-stain a blot with stains such as Coomassie or Sypro Ruby for total protein measurement. It takes just a few seconds to acquire a blot image and about 5 min to do total protein normalization using Image Lab software.
Second, the V3 workflow allows scientists to take better control of the western procedure because it makes the procedure more transparent and introduces several checkpoints for quality control. With the help of stain-free technology, researchers can visualize their protein samples on both the gel and the blot. Scientists can assess protein sample integrity (degraded or not), separation quality (precipitated or not), transfer efficiency and transfer quality (even transfer or not). These checkpoints help researches terminate the experiment when seeing big flaws in the process and avoid wasting time on poor samples and blots. This technology also helps scientists assess whether there is a significant amount of protein loss after membrane stripping and whether the blot is suitable for reprobing a different target 4.
Here are a few tips to ensure good experience and quality data using V3 western workflow.
<ol>
	<li>Image the gel immediately after gel run. Do not soak the gel in any buffer before the first imaging in the procedure as it may wash away the stain compound.</li>
	<li>Use the same imaging area to acquire all images in the protocol. This will allow the software to overlay images for data analysis such as total protein normalization.</li>
	<li>Keep the exposure time consistent when imaging the pretransfer and post-transfer gels. This will allow the software to quantitatively measure the transfer efficiency.</li>
	<li>Keep the membrane wet with a few drops of water or TBST when acquiring the blot image. This will avoid possible dirty background problems for target protein detection.</li>
	<li>Use low-fluorescent PVDF membrane for multiplexing fluorescent western blotting.</li>
</ol>
It is important to note that the stain-free molecule after UV-activation is irreversibly bound to tryptophan residues. This irreversible modification may potentially affect antigen recognition when using monoclonal antibodies if the epitope contains tryptophan. Polyclonal antibodies are unlikely to be affected because they recognize multiple epitopes on the antigen. Unbound stain-free molecules are easily washed off the gel and the membrane and therefore will not interfere with antibody-antigen interactions.
In conclusion, the V3 western workflow makes the western blot process faster, more transparent, quantitative and more reliable. Researchers now can easily apply total protein loading control in a western blot experiment to make their data more trustworthy. The V3 stain-free workflow has been adopted by a number of labs, and their publications have demonstrated that journals accept stain-free data as loading control in western blot22,17,7,8,5,23,18,15.

Western blot is a very useful technique9, however, there are two major challenges with western blotting: long and labor intensive process and quality of data. A traditional protocol requires about 2 days. It involves many steps including sample preparation, gel casting, protein electrophoresis and transfer, membrane blocking followed by antibody incubation, imaging, and quite often stripping, reprobing, and finally data analysis. Throughout this process, there are no reliable and flexible tools for process control. As such, errors can be introduced at each step, and these errors have the potential to generate data artifacts; therefore, loading controls are essential in western blotting to identify and correct for the errors. The loading control is usually done by checking the protein level of a reference protein in each sample to see if it is equally presented. People often use housekeeping proteins, such as &#946;-actin, &#946;-tubulin, GAPDH, as loading control.
The quality of western blot data depends on reliable loading control. But there are two legitimate concerns when using housekeeping proteins for loading controls: 1) the antibody-based immunodetection of the housekeeping protein bands are often saturated and therefore one cannot distinguish the loading differences among the samples30; 2) the housekeeping protein expression level may vary in the samples under certain experimental conditions , for example, siRNA treatment, cell death, cell differentiation,&#160;etc.11,28,3,6,10,21. Due to these concerns, scientific journals are now requiring that &#34;for quantitative comparisons, appropriate reagents, controls and imaging methods with linear signal ranges should be used&#34; (Nature guideline). Similarly, editors from the Journal of Clinical Investigation are asking for more reliable loading controls24. For these reasons, a housekeeping protein needs to be validated in order to be used as loading control. First, one has to make sure it is measured in the linear dynamic range of the immunodetection method14,29. Second, one has to make sure it is expressed consistently in all samples26,31,25,19,20.
An alternative solution to a reliable loading control is to use total protein measurement from the blot. Some researchers have stained the blots with total protein stains, such as Coomassie, Flamingo Pink, Sypro Ruby, Amido Black, Ponceau S, and stain-free technology, to measure the total protein signal in each lane as loading control16,20,13,27,1,4,12. The total protein loading control avoids the pitfalls associated with housekeeping proteins. First, it is a true reflection of the amount of protein loaded for each sample. Second, the total protein stain exhibits excellent linear dynamic range in the common loading range for western blot analysis (10-50 &#956;g protein of a complex cell lysate) and accurately differentiates the loading difference among the samples12.
Stain-free technology is a novel total protein staining method where a unique compound is mixed in acrylamide gel solution and evenly distributed in the casted gel. After electrophoresis is completed, the gel is exposed to UV light for a minimum of 1 min so that the stain compound reacts with the tryptophan residues in the protein. The proteins become excitable under UV light to give a strong fluorescent signal that can be visualized and quantified in a stain-free enabled imager such as the ChemiDoc MP system. The stain-free compound itself, however, does not absorb UV light, resulting in low background of the gel image. The modification of the tryptophan residues is irreversible and proteins can be visualized not only in the gel but also on the blot at any time after protein transfer.
The stain-free technology is applied in the V3 Western Workflow (Figure 1) to address the major complaints about the traditional workflow, especially the concerns with using housekeeping proteins as loading controls. Using this workflow, one could: 1) run a gel in about 20-30 min, 2) check sample integrity and protein separation quality in 5 min after gel run; 3) transfer proteins in 3-10 min; 4) check the transfer efficiency quantitatively; and 5) most importantly, validate changes in the level of the protein of interest using total protein loading control.

<table><tbody><tr><td colspan="4">&lt;strong&gt;REAGENTS&lt;/strong&gt;</td></tr><tr><td>4-20% Criterion TGX Stain-Free Precast Gel</td><td>Bio-Rad</td><td>&lt;strong&gt;567-8093&lt;/strong&gt;</td><td>Choose a gel size and percentage that meet the specific need in a experiment</td></tr><tr><td>Trans-Blot Turbo Midi PVDF Transfer Packs</td><td>Bio-Rad</td><td>&lt;strong&gt;170-4157&lt;/strong&gt;</td><td>Choose either PVDF or nitrocellulose and match the membrane size to the gel size</td></tr><tr><td>Clarity Western ECL Substrate, 500 ml </td><td>Bio-Rad</td><td>170-5061</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;EQUIPMENT&lt;/strong&gt;</td></tr><tr><td>Criterion Cell </td><td>Bio-Rad</td><td>165-6100</td><td></td></tr><tr><td>Trans-Blot Turbo Transfer Starter System </td><td>Bio-Rad</td><td>170-4155</td><td></td></tr><tr><td>ChemiDoc MP System </td><td>Bio-Rad</td><td>170-8280</td><td></td></tr></tbody></table>

v3 stain-free workflow for a practical, convenient, and reliable total protein loading control in western blotting

The western blot is a very useful and widely adopted lab technique, but its execution is challenging. The workflow is often characterized as a &#34;black box&#34; because an experimentalist does not know if it has been performed successfully until the last of several steps. Moreover, the quality of western blot data is sometimes challenged due to a lack of effective quality control tools in place throughout the western blotting process. Here we describe the V3 western workflow, which applies stain-free technology to address the major concerns associated with the traditional western blot protocol. This workflow allows researchers: 1) to run a gel in about 20-30 min; 2) to visualize sample separation quality within 5 min after the gel run; 3) to transfer proteins in 3-10 min; 4) to verify transfer efficiency quantitatively; and most importantly 5) to validate changes in the level of the protein of interest using total protein loading control. This novel approach eliminates the need of stripping and reprobing the blot for housekeeping proteins such as &#946;-actin, &#946;-tubulin, GAPDH, etc. The V3 stain-free workflow makes the western blot process faster, transparent, more quantitative&#160;and reliable.

1. Assessment of sample integrity, protein separation quality, and transfer efficiency with stain-free gel images.
Protein extract from HeLa cells were separated at 300 V for 20 min on a 18-well Criterion AnyKD TGX stain-free gel. The protein samples were loaded 3x at four different amounts (Lanes 1-3, 40 &#956;g; Lanes 4-6, 30 &#956;g; Lanes 7-9, 20 &#956;g; Lanes 10-12, 10 &#956;g). The gel was activated under UV light for 1 min. Figure 2A shows the gel image acquired right after the protein separation. Protein sample integrity (e.g. degradation) and separation quality (e.g. protein precipitation) can be visually assessed with this gel image. Proteins were then transferred for 7 min to a nitrocellulose membrane using Trans-Blot Turbo. Figure 2B shows the stain-free image of the post-transfer gel. Both images were acquired with the same exposure time (6.8 sec). Lane 3 and 12 were selected to measure the transfer efficiency. Using the volume tool in the Image Lab software, a rectangular box (blue) was drew to cover lane 3 and 12 on both gel images. The calculation based on the volume values from these boxes indicated that transfer efficiency of both lanes was 80% (Figure 2C). In this experiment, the AnyKD TGX gel was selected to study small to medium size target proteins and it was not optimized for the transfer of large proteins. Optimizing the transfer efficiency would require the use of lower percentage gel (e.g. 4-20%) and/or an adjustment of the transfer time to facilitate the transfer of large proteins. 2. Stain-free total protein loading control is a reliable alternative to housekeeping loading control in western blotting to quantify a small change in the level of protein of interest.
MCM-7 is a DNA licensing replication factor the level of which decreases by 20-50% in Lymphoblastoid cell lines (LCL) after irradiation treatment. In this experiment, lysates (30 &#956;g each) of four control and irradiation-treated Lymphoblastoid cell line (LCL) cultures were separated on a 12-well Criterion AnyKD TGX stain-free gel. The gel was activated for 1 min under UV light and transferred by Trans-Blot Turbo to a PVDF membrane for immunoblotting. The housekeeping protein GAPDH (green) was probed with a rabbit antibody (Cell Signaling Technology, USA, 1:2,500) and a Dylight 549 conjugated Goat-anti-rabbit antibody (Rockland, USA, 1:20,000). The protein of interest MCM-7 (red) was probed using a mouse antibody (Abcam, USA, 1:1,000) and a Dylight 649 conjugated Goat-anti-mouse antibody (Rockland, 1:10,000).
Figure 3A shows a multiplex fluorescent image of total proteins (blue), MCM-7 (red) and GAPDH (green) detected in four control and irradiation treated LCL samples. Figure 3B is a stain-free image of the same blot showing the total protein patterns in each sample (30 &#956;g). Image lab software selected the sample lanes (blue boxes) to measure MCM-7, GAPDH, and total protein volume in each lane. The MCM-7 levels were normalized either against the stain-free total protein measurement or against GAPDH. The normalized MCM-7 protein levels were statistically analyzed and the average MCM-7 protein band volume and standard deviation (n=4) are presented in the chart (Figure 3C). Both normalization methods revealed a small decrease (about 25%) in MCM-7 protein levels after irradiation treatment. The data with the total protein normalization exhibited a smaller standard deviation than that with GAPDH as the loading control.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50948/50948fig1highres.jpg" src="/files/ftp_upload/50948/50948fig1.jpg" /> 
Figure 1. V3 Western Workflow. The V3 workflow is depicted in the left column in 4 steps. The major instruments and reagents used in the workflow are shown at each step. The estimated time for each step is also included. The right column shows that a minimum of 4 images can be generated in the V3 workflow. The use of each piece of data is described. The stain-free images of the pretransfer gel, post-transfer gel, and the blot (A, B, C) can not be generated easily with traditional western blotting techniques; These images and data provide important information and checkpoints along the procedure to improve the scientist&#39;s control and reproducibility of western blot workflow. The target protein signals can be captured either on a chemiluminescent blot image (D) if a HRP-conjugated secondary antibody was applied in detection or on a fluorescent blot image (E) if multiplexing fluorescent western blotting was performed to detect more than one target protein simultaneously on the same blot. 
<a href="https://www.jove.com/files/ftp_upload/50948/50948fig1highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50948/50948fig2highres.jpg" src="/files/ftp_upload/50948/50948fig2.jpg" /> 
Figure 2. Stain-free images of pretransfer and post-transfer gels in the V3 western workflow to assess sample integrity, separation quality, and transfer efficiency. (A) Pretransfer gel stain-free image. (B) Post-transfer gel stain-free image. (C) Protein transfer efficiency measurement.
<a href="https://www.jove.com/files/ftp_upload/50948/50948fig2highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 3" fo:content-width="3.5in" fo:src="/files/ftp_upload/50948/50948fig3highres.jpg" src="/files/ftp_upload/50948/50948fig3.jpg" /> 
Figure 3. Comparing stain-free total protein measurement to GAPDH immunodetection as loading controls to normalize the level of protein of interest. (A) Fluorescent western blot image. (B) Stain-free blot image. (C) Normalized MCM-7 protein levels. 
<a href="https://www.jove.com/files/ftp_upload/50948/50948fig3highres.jpg" target="_blank">Click here to view larger image.</a>

Watch this Scientific Journal Video about V3 Stain-free Workflow for a Practical, Convenient, and Reliable Total Protein Loading Control in Western Blotting at JoVE.com

V3 Stain-free Workflow for a Practical, Convenient, and Reliable Total Protein Loading Control in Western Blotting

V3 workflow is a western blot procedure using stain-free gels. The stain-free technology allows researchers to visualize protein separation quality, to verify the transfer efficiency, and most importantly, to validate the change in the protein of interest using total protein quantification as a reliable loading control.

The western blot is a very useful and widely adopted lab technique, but its execution is challenging. The workflow is often characterized as a &#34;black ...

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22.5K Views.  Bio-Rad Laboratories. V3 workflow is a western blot procedure using stain-free gels. The stain-free technology allows researchers to visualize protein separation quality, to verify the transfer efficiency, and most importantly, to validate the change in the protein of interest using total protein quantification as a reliable loading control.

Video: V3 Stain-free Workflow for a Practical, Convenient, and Reliable Total Protein Loading Control in Western Blotting

Western Blotting Using the Invitrogen NuPage Novex Bis Tris MiniGels

Western Blotting (or immunoblotting) is a standard laboratory procedure allowing investigators to verify the expression of a protein, determine the relative amount of the protein present in different samples, and analyze the results of co-immunoprecipitation experiments. In this method, a target protein is detected with a specific primary antibody in a given sample of tissue homogenate or extract. Protein separation according to molecular weight is achieved using denaturing SDS-PAGE. After transfer to a membrane, the target protein is probed with a specific primary antibody and detected by chemiluminescence.
Since its first description, the western-blotting technique has undergone several improvements, including pre-cast gels and user-friendly equipment. In our laboratory, we have chosen to use the commercially available NuPAGE electrophoresis system from Invitrogen. It is an innovative neutral pH, discontinuous SDS-PAGE, pre-cast mini-gel system. This system presents several advantages over the traditional Laemmli technique including: i) a longer shelf life of the pre-cast gels ranging from 8 months to 1 year; ii) a broad separation range of molecular weights from 1 to 400 kDa depending of the type of gel used; and iii) greater versatility (range of acrylamide percentage, the type of gel, and the ionic composition of the running buffer).
The procedure described in this video article utilizes the Bis-Tris discontinuous buffer system with 4-12% Bis-Tris gradient gels and MES running buffer, as an illustration of how to perform a western-blot using the Invitrogen NuPAGE electrophoresis system. In our laboratory, we have obtained good and reproducible results for various biochemical applications using this western-blotting method.

This technical article describes a standard western-blotting procedure using the commercially available NuPAGE electrophoresis Mini-Gel system from Invitrogen.

Western Blotting (or immunoblotting) is a standard laboratory procedure allowing investigators to verify the expression of a protein, determine the ...

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 ...

Biochemistry

Robust Comparison of Protein Levels Across Tissues and Throughout Development Using Standardized Quantitative Western Blotting

Western blotting is a technique that is commonly used to detect and quantify protein expression. Over the years, this technique has led to many advances in both basic and clinical research. However, as with many similar experimental techniques, the outcome of Western blot analyses is easily influenced by choices made in the design and execution of the experiment. Specific housekeeping proteins have traditionally been used to normalize protein levels for quantification, however, these have a number of limitations and have therefore been increasingly criticized over the past few years. Here, we describe a detailed protocol that we have developed to allow us to undertake complex comparisons of protein expression variation across different tissues, mouse models (including disease models), and developmental timepoints. By using a fluorescent total protein stain and introducing the use of an internal loading standard, it is possible to overcome existing limitations in the number of samples that can be compared within experiments and systematically compare protein levels across a range of experimental conditions. This approach expands the use of traditional western blot techniques, thereby allowing researchers to better explore protein expression across different tissues and samples.

This method describes a robust and reproducible approach for the comparison of protein levels in different tissues and at different developmental timepoints using a standardized quantitative western blotting approach.

Western blotting is a technique that is commonly used to detect and quantify protein expression. Over the years, this technique has led to many advances ...

Western Blotting: Sample Preparation to Detection

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein.

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract.

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. It uses gel ...

The Fastest Western in Town: A Contemporary Twist on the Classic Western Blot Analysis

The Western blot techniques that were originally established in the late 1970s are still actively utilized today. However, this traditional method of Western blotting has several drawbacks that include low quality resolution, spurious bands, decreased sensitivity, and poor protein integrity. Recent advances have drastically improved numerous aspects of the standard Western blot protocol to produce higher qualitative and quantitative data. The Bis-Tris gel system, an alternative to the conventional Laemmli system, generates better protein separation and resolution, maintains protein integrity, and reduces electrophoresis to a 35 min run time. Moreover, the iBlot dry blotting system, dramatically improves the efficacy and speed of protein transfer to the membrane in 7 min, which is in contrast to the traditional protein transfer methods that are often more inefficient with lengthy transfer times. In combination with these highly innovative modifications, protein detection using infrared fluorescent imaging results in higher-quality, more accurate and consistent data compared to the standard Western blotting technique of chemiluminescence. This technology can simultaneously detect two different antigens on the same membrane by utilizing two-color near-infrared dyes that are visualized in different fluorescent channels. Furthermore, the linearity and broad dynamic range of fluorescent imaging allows for the precise quantification of both strong and weak protein bands. Thus, this protocol describes the key improvements to the classic Western blotting method, in which these advancements significantly increase the quality of data while greatly reducing the performance time of this experiment.

This protocol explores the latest advancements in performing Western blot analyses.&#160;These novel modifications employ a Bis-Tris gel system with a 35&#160;min electrophoresis run time, a 7&#160;min dry blotting transfer system, and infrared fluorescent protein detection and imaging that generates higher resolution, quality, sensitivity, and improved accuracy of Western data.

The Western blot techniques that were originally established in the late 1970s are still actively utilized today. However, this traditional method of ...

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...

Immunoprecipitation-Based Techniques: Purification of Endogenous Proteins Using Agarose Beads

Source: Susannah C. Shissler1, Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland, Baltimore, MD 21201
Immunoprecipitation (IP, also known as a &#39;pull-down&#39; assay) is a widely used technique that has applications in a variety of fields. First conceived in 1984, it was refined in 1988 (1, 2). The fundamental goal of IP is purification and isolation of a specific protein using an antibody against that protein. The word &#34;immuno&#34; refers to the use of an antibody while the word &#34;precipitation&#34; refers to pulling down a specific substance from a solution. The target protein might be endogenous or recombinant. Most recombinant proteins have an epitope tag (i.e. myc or flag) attached to them to simplify subsequent purification. Typically, it is easier to optimize recombinant protein IP because the antibodies against recombinant epitope tags are very strong and effective. Antibodies against endogenous proteins have extremely variable efficacy - making it much more difficult to optimize these IPs. A necessary step after immunoprecipitation is verification of purification. The isolated protein is resolved using SDS-PAGE and subsequently probed for purity by western blots (Figure 1). An important control is the use of a different antibody during the Western blot to verify pull down of the correct protein. The combination of IP with subsequent techniques is a powerful analysis tool. The goal after purification may be characterization of the protein itself by NMR, mass spectrometry, and in vitro assays, or analysis of the protein&#39;s interacting partners (i.e. protein, DNA, RNA) (3, 4, 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10502/10502fig1.jpg" /> 
Figure 1: Overview of Immunoprecipitation Procedure. Immunoprecipitation is the isolation of a specific protein using an antibody. After production of lysate from cells, there are two major steps- pre-clearing and pull down. During pre-clearing step, the cell-lysates are pre-cleared of proteins that bind to antibodies non-specifically using an isotype control antibody. In pull down step, the target protein is pulled down using a protein-specific antibody. The isolated protein is then analyzed by Western blot. Isotype antibodies and protein specific antibodies have the same constant domain, but different antigen binding domains. A key component of this protocol is Protein A/G agarose beads that bind the constant domain of antibodies- allowing immunoprecipitation of the target protein. <a href="https://www.jove.com/files/ftp_upload/10502/10502fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Antibodies are the key component of an immunoprecipitation that differentiate it from other forms of protein purification (i.e. nickel affinity column purification). Antibodies are molecules made by B cells that can recognize specific protein epitopes. Antibodies have two domains: constant (Fc) and antigen binding (Fab) (Figure 1). The constant domain identifies the type of antibody and dictates function in vivo. Usually, the constant domains of antibodies used for IP are mouse, rat, or rabbit IgG. The antigen binding portion of the antibody recognizes a specific epitope of a specific protein. Antibodies can recognize epitopes on folded proteins that may not exist when the protein is denatured and vice versa. Therefore, the availability of the epitope depends on protein folding - identifying an important factor to consider when choosing antibodies and conditions for IP.
Both prokaryotic and eukaryotic systems have antibody-binding proteins. In eukaryotic systems, the purpose is immune protection from bacteria while in prokaryotic systems, the purpose is protection from the immune system. Antibody-binding proteins affect IP methodology in two ways. First, there is a necessary pre-clearing step (Figure 1) to rid the lysate of proteins that bind antibodies - thereby reducing non-specific binding in the final product. This step uses an isotype antibody that has the same constant domain as but a different antibody binding domain than your protein-specific antibody. Bacterial antibody-binding proteins are the second key component of this method. After the protein-specific antibody binds the target protein, the antibody: protein complex must be pulled down (Figure 1). Proteins A, G, and L are bacterial proteins that bind the constant domain of antibodies. While bacteria use this to subvert the immune system, researchers have co-opted this system for easy antibody purification, and it is used during both the pre-clearing and pull-down steps. These proteins have different binding affinities for different species and different constant domain subtypes - another factor to consider when choosing conditions for IP. Many companies sell Protein A/G labeled agarose beads (Figure 1), pre-made spin columns, or resins to make columns. In general, beads and spin columns are used for smaller sample sizes while resins are used for bulk purification.
In this lab exercise, we demonstrate how to purify the endogenous protein c-myc, from primary murine thymocytes, using Protein A/G Plus agarose beads based basic immunoprecipitation technique. The protocol starts from cell lysate preparation and ends with the with verification of successful protein pull down using Western blot analysis.

Immunoprecipitation Techniques: Purification of Proteins

Source: Susannah C. Shissler1, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland, Baltimore, MD 21201
Immunoprecipitation ...

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The Western Blot

Western Blotting is used to identify the presence of specific proteins in electrophoretically separated samples. Following separation by a technique known as sodium dodecyl sulfate polyacrylamide gel electrophoresis, or SDS-PAGE, western transfer is used to move proteins from a polyacrylamide gel onto a piece of membrane which traps the proteins in their respective locations. Next, the membranes are probed with antibodies in a process called immunboblotting. Immunoblotting uses antibody-protein and antibody-antibody binding through specific recognition sites, providing the high specificity required for identifying a single protein. The detection of antibodies takes place using reporter systems which includes the use of enzymes. Enzymes can be attached to the end of an antibody and react with substrates to produce changes in color or light. These signals can then be imaged and quantified using a process called densitometry.
This video-article presents an overview of the western blot technique by describing western transfer, the use of antibody detection, and image analysis. The steps involved with western transfer such as the assembly of the transfer sandwich and transfer conditions are discussed in detail as well as the theory behind antibody binding and detection of those antibodies. The broad applications of this technique are described through several examples including the detection of protein-protein interactions and identification of individual proteins within protein complexes.

Western Blotting: Western Transfer, Antibody Detection, and Image Analysis

Western Blotting is used to identify the presence of specific proteins in electrophoretically separated samples.  Following separation by a technique ...

Basic Biology

Basic Methods in Cellular and Molecular Biology

Quantitative Western Blotting to Estimate Protein Expression Levels

Source: Huang, Y. T., et al. <a href="https://app.jove.com/v/59438">Robust Comparison of Protein Levels Across Tissues and Throughout Development Using Standardized Quantitative Western Blotting.</a> J. Vis. Exp. (2019).
The video demonstrates a total protein normalization-based western blotting technique for quantifying the target protein expression in a sample. By utilizing the total protein staining of the sample, a ratiometric analysis is performed to determine the relative expression level of the target protein.