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 border="1">
 <tbody>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>REAGENTS</td>
 </tr>
 <tr>
 <td>4-20% Criterion TGX Stain-Free Precast Gel</td>
 <td>Bio-Rad</td>
 <td>567-8093 </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>170-4157 </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></td>
 <td></td>
 <td></td>
 <td>[header]</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>EQUIPMENT</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 vistas.  Bio-Rad Laboratories. El flujo de trabajo V3 es un procedimiento de western blot que utiliza geles sin manchas. La tecnología libre de manchas permite a los investigadores visualizar la calidad de la separación de proteínas, verificar la eficiencia de la transferencia y, lo que es más importante, validar el cambio en la proteína de interés utilizando la cuantificación total de proteínas como un control de carga confiable.

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 ...

The Ladder Rung Walking Task: A Scoring System and its Practical Application.

Progress in the development of animal models for/stroke, spinal cord injury, and other neurodegenerative disease requires tests of high sensitivity to elaborate distinct aspects of motor function and to determine even subtle loss of movement capacity. To enhance efficacy and resolution of testing, tests should permit qualitative and quantitative measures of motor function and be sensitive to changes in performance during recovery periods. The present study describes a new task to assess skilled walking in the rat to measure both forelimb and hindlimb function at the same time. Animals are required to walk along a horizontal ladder on which the spacing of the rungs is variable and is periodically changed. Changes in rung spacing prevent animals from learning the absolute and relative location of the rungs and so minimize the ability of the animals to compensate for impairments through learning. In addition, changing the spacing between the rungs allows the test to be used repeatedly in long-term studies. Methods are described for both quantitative and qualitative description of both fore- and hindlimb performance, including limb placing, stepping, co-ordination. Furthermore, use of compensatory strategies is indicated by missteps or compensatory steps in response to another limb&#x2019;s misplacement. 

The ladder rung walking task is a new test to assess skilled walking and measure both forelimb and hindlimb placing, stepping, and inter-limb co-ordination.

Progress in the development of animal models for/stroke, spinal cord injury, and other neurodegenerative disease requires tests of high sensitivity to ...

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 ...

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation (BiFC) System

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein (YFP) into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest. These constructs can then be co-transformed into Nicotiana benthamiana via Agrobacterium mediated transformation, allowing the transit expression of fusion proteins. The reconstitution of YFP signal only occurs when the inquest proteins interact 1-7. To test and validate the protein-protein interactions, BiFC can be used together with yeast two hybrid (Y2H) assay. This may detect indirect interactions which can be overlooked in the Y2H. Gateway technology is a universal platform that enables researchers to shuttle the gene of interest (GOI) into as many expression and functional analysis systems as possible8,9. Both the orientation and reading frame can be maintained without using restriction enzymes or ligation to make expression-ready clones. As a result, one can eliminate all the re-sequencing steps to ensure consistent results throughout the experiments. We have created a series of Gateway compatible BiFC and Y2H vectors which provide researchers with easy-to-use tools to perform both BiFC and Y2H assays10. Here, we demonstrate the ease of using our BiFC system to test protein-protein interactions in N. benthamiana plants.

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation BiFC System

We have developed a technique to test protein-protein interactions in plant. A yellow fluorescent protein (YFP) is split into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest via Gateway system, enabling expression of fusion proteins. Reconstitution of YFP signal only occurs when the inquest proteins interact.

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein ...

A Quantitative Fitness Analysis Workflow

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. QFA can be applied to focused observations of single cultures but is most useful for genome-wide genetic interaction or drug screens investigating up to thousands of independent cultures. The central experimental method is the inoculation of independent, dilute liquid microbial cultures onto solid agar plates which are incubated and regularly photographed. Photographs from each time-point are analyzed, producing quantitative cell density estimates, which are used to construct growth curves, allowing quantitative fitness measures to be derived. Culture fitnesses can be compared to quantify and rank genetic interaction strengths or drug sensitivities. The effect on culture fitness of any treatments added into substrate agar (e.g. small molecules, antibiotics or nutrients) or applied to plates externally (e.g. UV irradiation, temperature) can be quantified by QFA.
The QFA workflow produces growth rate estimates analogous to those obtained by spectrophotometric measurement of parallel liquid cultures in 96-well or 200-well plate readers. Importantly, QFA has significantly higher throughput compared with such methods. QFA cultures grow on a solid agar surface and are therefore well aerated during growth without the need for stirring or shaking.
QFA throughput is not as high as that of some Synthetic Genetic Array (SGA) screening methods5,6. However, since QFA cultures are heavily diluted before being inoculated onto agar, QFA can capture more complete growth curves, including exponential and saturation phases3. For example, growth curve observations allow culture doubling times to be estimated directly with high precision, as discussed previously1.
Here we present a specific QFA protocol applied to thousands of S. cerevisiae cultures which are automatically handled by robots during inoculation, incubation and imaging. Any of these automated steps can be replaced by an equivalent, manual procedure, with an associated reduction in throughput, and we also present a lower throughput manual protocol. The same QFA software tools can be applied to images captured in either workflow.
We have extensive experience applying QFA to cultures of the budding yeast S. cerevisiae but we expect that QFA will prove equally useful for examining cultures of the fission yeast S. pombe and bacterial cultures.

Quantitative Fitness Analysis (QFA) is a complementary series of experimental and computational methods for estimating microbial culture fitnesses. QFA estimates the effect of genetic mutations, drugs or other applied treatments on microbe growth. Experiments scaling from focussed analysis of single cultures to thousands of parallel cultures can be designed.

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. ...

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and behavioural research. As a vertebrate, zebrafish share considerable genetic sequence similarity with humans and are being used as an animal model for various human disease conditions. The advantages of zebrafish in comparison to other common vertebrate models include high fecundity, low maintenance cost, transparent embryos, and rapid development. Due to the spur of interest in zebrafish research, the need to establish and maintain a productive zebrafish housing facility is also increasing. Although literature is available for the maintenance of a zebrafish laboratory, a concise video protocol is lacking. This video illustrates the protocol for regular housing, feeding, breeding and raising of zebrafish larvae. This process will help researchers to understand the natural behaviour and optimal conditions of zebrafish husbandry and hence troubleshoot experimental issues that originate from the fish husbandry conditions. This protocol will be of immense help to researchers planning to establish a zebrafish laboratory, and also to graduate students who are intending to use zebrafish as an animal model.

Regular Care and Maintenance of a Zebrafish Danio rerio Laboratory: An Introduction

This protocol outlines regular maintenance and care to maintain optimal conditions for zebrafish husbandry. The video illustrates the protocol for system maintenance, regular housing, feeding, breeding, and raising of zebrafish larvae.

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and ...

A Practical and Novel Method to Extract Genomic DNA from Blood Collection Kits for Plasma Protein Preservation

Laboratory tests can be done on the cellular or fluid portions of the blood. The use of different blood collection tubes determines the portion of the blood that can be analyzed (whole blood, plasma or serum). Laboratories involved in studying the genetic basis of human disorders rely on anticoagulated whole blood collected in EDTA-containing vacutainer as the source of DNA for genetic / genomic analysis. Because most clinical laboratories perform biochemical, serologic and viral testing as a first step in phenotypic outcome investigation, anticoagulated blood is also collected in heparin-containing tube (plasma tube). Therefore when DNA and plasma are needed for simultaneous and parallel analyses of both genomic and proteomic data, it is customary to collect blood in both EDTA and heparin tubes. If blood could be collected in a single tube and serve as a source for both plasma and DNA, that method would be considered an advancement to existing methods. The use of the compacted blood after plasma extraction represents an alternative source for genomic DNA, thus minimizing the amount of blood samples processed and reducing the number of samples required from each patient. This would ultimately save time and resources.
The BD P100 blood collection system for plasma protein preservation were created as an improved method over previous plasma or serum collection tubes1, to stabilize the protein content of blood, enabling better protein biomarker discovery and proteomics experimentation from human blood. The BD P100 tubes contain 15.8 ml of spray-dried K2EDTA and a lyophilized proprietary broad spectrum cocktail of protease inhibitors to prevent coagulation and stabilize the plasma proteins. They also include a mechanical separator, which provides a physical barrier between plasma and cell pellets after centrifugation. Few methods have been devised to extract DNA from clotted blood samples collected in old plasma tubes2-4. Challenges from these methods were mainly associated with the type of separator inside the tubes (gel separator) and included difficulty in recovering the clotted blood, the inconvenience of fragmenting or dispersing the clot, and obstruction of the clot extraction by the separation gel.
We present the first method that extracts and purifies genomic DNA from blood drawn in the new BD P100 tubes. We compare the quality of the DNA sample from P100 tubes to that from EDTA tubes. Our approach is simple and efficient. It involves four major steps as follows: 1) the use of a plasma BD P100 (BD Diagnostics, Sparks, MD, USA) tube with mechanical separator for blood collection, 2) the removal of the mechanical separator using a combination of sucrose and a sterile paperclip metallic hook, 3) the separation of the buffy coat layer containing the white cells and 4) the isolation of the genomic DNA from the buffy coat using a regular commercial DNA extraction kit or a similar standard protocol.

We are describing a new method of isolating genomic DNA from whole blood collected for plasma/serology. After plasma collection, the compacted blood is usually discarded. Our novel method represents a significant improvement over existing methods and makes DNA and plasma available from a single collection, without requesting additional blood.

Laboratory tests can be done on the cellular or  fluid portions of the blood. The use of different blood collection tubes  determines the portion of the ...

A Practical Guide to Phylogenetics for Nonexperts

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this topic and so it presents inherent problems. Here we compile a practical introduction to phylogenetics for nonexperts.&#160;We outline in a step-by-step manner, a pipeline for generating reliable phylogenies from gene sequence datasets. We begin with a user-guide for similarity search tools via online interfaces as well as local executables. Next, we explore programs for generating multiple sequence alignments followed by protocols for using software to determine best-fit models of evolution. We then outline protocols for reconstructing phylogenetic relationships via maximum likelihood and Bayesian criteria&#160;and finally describe tools for visualizing phylogenetic trees. While this is not by any means an exhaustive description of phylogenetic approaches, it does provide the reader with practical starting information on key software applications commonly utilized by phylogeneticists. The vision for this article would be that it could serve as a practical training tool for researchers embarking on phylogenetic studies&#160;and also serve as an educational resource that could be incorporated into a classroom or teaching-lab.

Here we describe a step-by-step pipeline for generating reliable phylogenies from nucleotide or amino acid sequence datasets. This guide aims to serve researchers or students new to phylogenetic analysis.&#160;

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this ...

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 ...

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live cells. BRET is the non-radiative transfer of energy from a 'donor' luciferase enzyme to an 'acceptor' fluorescent protein. In the most common configuration of this assay, the donor is Renilla reniformis luciferase and the acceptor is Yellow Fluorescent Protein (YFP). Because the efficiency of energy transfer is strongly distance-dependent, observation of the BRET phenomenon requires that the donor and acceptor be in close proximity. To test for an interaction between two proteins of interest in cultured mammalian cells, one protein is expressed as a fusion with luciferase and the second as a fusion with YFP. An interaction between the two proteins of interest may bring the donor and acceptor sufficiently close for energy transfer to occur. Compared to other techniques for investigating protein-protein interactions, the BRET assay is sensitive, requires little hands-on time and few reagents, and is able to detect interactions which are weak, transient, or dependent on the biochemical environment found within a live cell. It is therefore an ideal approach for confirming putative interactions suggested by yeast two-hybrid or mass spectrometry proteomics studies, and in addition it is well-suited for mapping interacting regions, assessing the effect of post-translational modifications on protein-protein interactions, and evaluating the impact of mutations identified in patient DNA.

Interactions between proteins are fundamental to all cellular processes. Using Bioluminescence Resonance Energy Transfer, the interaction between a pair of proteins can be monitored in live cells and in real time. Furthermore, the effects of potentially pathogenic mutations can be assessed.

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live ...