E.J.N.G. is supported by the Wellcome Trust (grant 106098/Z/14/Z). Other funding has been provided by the SMA Trust (SMA UK Consortium; T.H.G. &#38; Y-T.H.), SMA Europe (T.H.G, D.v.D.H. &#38; E.J.N.G.), the University of Edinburgh DTP in Precision Medicine (T.H.G., L.L. &#38; A.M.M.), and the Euan MacDonald Centre for Motor Neurone Disease Research (T.H.G).

Tissues for this procedure were obtained from animal studies that were approved by the internal ethics committee at the University of Edinburgh and were performed in concordance with institutional and UK Home Office regulations under the authority of relevant personal and project licenses.
NOTE: This protocol has been optimized using standardized, commercially available kits and reagents in order to increase reproducibility (see Table of Materials).
1. Preparation of samples
<ol>
	<li>Protein extraction 
	<ol>
		<li>Transfer snap-frozen cell or tissue samples from -80 &#176;C on dry ice, thaw on ice, and wash as required with ice cold 1x phosphate-buffered saline (PBS) (for details on tissues and PBS washes, see Table 1). Avoid unnecessary freeze-thaw cycles as this will affect protein quality.</li>
		<li>Add radioimmunoprecipitation assay (RIPA) buffer (25 mM Tris-HCl (pH 7.6), 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS) containing 1x protease inhibitor to each sample, using the optimal amount per tissue weight (see Table 1 for recommendations). 
		NOTE: Depending on the application, the type and amount of homogenization buffer may need further optimization.</li>
		<li>Use a hand-held electric homogenizer with a polypropylene pestle to homogenize tissue samples. Between each sample, wash the pestle in double-distilled water and dry with a clean tissue. Change the pestle between different experimental conditions and tissues.</li>
		<li>Leave the samples on ice for 10 min after homogenization. Centrifuge the samples at &#62;10,000 x g at 4 &#176;C for 10 min.</li>
		<li>Transfer the supernatant to a new tube on ice without disturbing the pellet. The supernatant is the protein sample. Store the extracted protein at -80 &#176;C or directly proceed to measuring the protein concentration.</li>
	</ol>
	</li>
	<li>Quantification and normalization of protein concentration 
	<ol>
		<li>Measure the protein concentration using a bicinchoninic acid (BCA) assay. Prepare a BCA assay mix according to the manufacturer&#8217;s instructions in a 96-well optical plate using 200 &#181;L of BCA mix per well. 
		NOTE: Other quantification methods such as Lowry and Bradford assays can also be used to determine protein concentration as long as protein concentration is quantified consistently across experiments.</li>
		<li>Prepare bovine serum albumin (BSA) standards at increasing concentrations in triplicate and add 1 &#181;L of each protein sample in duplicate. Incubate the 96-well plate in a preheated heat block at 60 &#176;C for 10 min or longer if the protein concentration is expected to be low.</li>
		<li>After incubation, measure the absorption at 560 nm using a plate reader.</li>
		<li>Export the plate reader measurements and calculate the protein concentration by comparing the average absorbance values of each sample to a standard curve obtained using the protein standard. The R-squared value for the standard curve should be greater than or equal to 0.98 to accurately estimate sample protein concentration.</li>
		<li>Normalize the amount of protein by preparing dilutions of protein samples in sample buffer and ultrapure water. The total volume can be adjusted depending on the type of gel used. Loading 30 &#181;g of protein per lane as a starting amount is recommended. Add reducing agent such as dithiothreitol (DTT; final concentration 5 mM) or beta-mercaptoethanol (final concentration 200 mM) to each sample as required. Pipette undiluted beta-mercaptoethanol in a fume hood.</li>
		<li>Incubate the samples in a heat block at 70 &#176;C for 10 min. Put the samples on ice, vortex and spin down briefly to collect. Keep on ice until loading the gel.</li>
	</ol>
	</li>
</ol>
2. Gel electrophoresis of protein samples
<ol>
	<li>Device and gel set up
	<ol>
		<li>Setup a precast 4&#8211;12% Bis-Tris gradient gel (see Table of Materials) in the gel electrophoresis chamber system. Rinse the gels using double-distilled water before use. 
		NOTE: Depending on the size, interactions and abundance of the protein of interest, gels with a different gradient, buffering agent or well size and number can be used.</li>
		<li>Add 500 mL of 1x MES SDS running buffer diluted in double-distilled water per tank. Carefully remove the comb from the gels after adding the running buffer without disturbing the wells in the stacking gel.</li>
	</ol>
	</li>
	<li>Protein loading
	<ol>
		<li>Load 3.5 &#181;L of a protein standard into the well. Depending on the sample layout, loading a protein ladder on both sides of the gel can aid in more accurately estimating protein size. Use fine-tipped gel loading tips for more accurate sample loading.</li>
		<li>When using an internal standard for between-membrane normalization (see step 5 below and discussion), load an amount that is equal to the other samples into the first 3 wells next to the protein ladder.</li>
		<li>Load 30 &#181;g of each sample in the remaining wells. Add 1x sample buffer to all empty wells.</li>
	</ol>
	</li>
	<li>Electrophoresis
	<ol>
		<li>Assemble the gel tank after loading the samples. Run the samples through the stacking gel at 80 V for 10 min followed by 150 V for an additional 45-60 min.</li>
	</ol>
	</li>
</ol>
3. Protein transfer
NOTE: Protein transfer in this protocol is performed using a commercially available semi-dry blotting system (see Table of Materials) for fast and consistent outcomes.
<ol>
	<li>Prepare the protein transfer by pre-soaking filter paper in double-distilled water and ensuring the gel knife, plastic Pasteur pipette, blotting roller and forceps are ready to use.</li>
	<li>Open the transfer stack by carefully removing all wrapping foil. Remove the top from the bottom stack and set it aside. Quickly moisten the membrane on the bottom stack with several drops of electrophoresis running buffer (2-3 mL). Once the transfer stack is open, it is important to prevent the PVDF membrane from drying out.</li>
	<li>After stopping the electrophoresis, open the pre-cast gel using the gel knife and cut off the stacking gel. Cut the gel around its edges to free it from the plastic cast. Keep the gel knife wet to prevent damage to the gel.</li>
	<li>Assemble the transfer stack from bottom to top: bottom stack (containing the PVDF membrane), protein gel, filter paper. Use the blotting roller to remove all air bubbles. Place the top stack on top of the filter paper and roll the stack again to remove air bubbles. Do not push too strongly as this may cause the gel to deform during protein transfer.</li>
	<li>Transfer the whole stack into the transfer device with the electrode on the left side of the device and place the gel sponge on top of the stack so that it is aligned with the corresponding electrical contacts on the device. Close the lid, select and start the appropriate program (20 V for 7 min is a recommended starting point).</li>
	<li>When finished, leave the lid closed for 2 min to allow the stack to cool down and to prevent the membrane from drying out. Remove the transfer stack and cut the membrane to the gel size. Wash the cut membrane quickly with double-distilled water before continuing with the total protein stain.</li>
</ol>
4. Total protein staining
NOTE: Using fluorescent detection provides a substantial benefit over more traditional approaches (e.g., ECL detection), as the linear range and sensitivity can be much better controlled4. Therefore, in steps 4 and 5, a fluorescent TPS and fluorescent secondary antibodies are used (see Table of Materials).
<ol>
	<li>Roll the membrane into a 50-mL tube with the protein side facing inwards. Because of light sensitivity of the fluorescent TPS and secondary antibodies, all subsequent steps are carried out in the dark.</li>
	<li>Add 5 mL of protein stain solution (see Table of Materials) and incubate on a roller for 5 min at room temperature. Because TPS and wash buffer contain methanol, carry these steps out in a fume hood.</li>
	<li>Discard the staining solution and wash twice quickly with the 5 mL of wash solution (6.3% acetic acid in 30% methanol). Place the tube briefly back on the roller between wash steps. Rinse the membrane briefly with ultrapure water before continuing.</li>
</ol>
5. Blocking, antibody incubation and detection
<ol>
	<li>Blocking the membrane: Add 3 mL of blocking buffer (see Table of Materials) to the 50 mL tube containing the membrane. Incubate the membrane on a roller for 30 min at room temperature. Depending on the choice of antibody, the type of blocking buffer used may require optimization.</li>
	<li>Primary antibody incubation
	<ol>
		<li>Discard the blocking buffer and replace with the primary antibody at the appropriate, optimized concentration (Figure 1 and Figure 2: mouse-anti-SMN, at 1:1,000, diluted in blocking buffer). 
		NOTE: Adequate optimization of primary antibodies should include confirmation that the antibody detects a protein product of the right size whilst showing no or minimal binding to other, unspecific protein products. If possible, test and compare multiple antibodies against the protein of interest.</li>
		<li>Incubate the membrane on a roller overnight at 4 &#176;C. The next day, remove the antibody solution and wash 6 times for 5 min with 1x PBS on a roller at room temperature (RT).</li>
	</ol>
	</li>
	<li>Secondary antibody incubation
	<ol>
		<li>Prepare the specific secondary antibody at 1:5,000 against the host of the primary antibody in 5 mL blocking buffer. Other secondary antibodies may require other dilutions or the use of alternative blocking buffers.</li>
		<li>Incubate the membrane with the secondary antibody solution on a roller for 1 h at RT. After incubation, wash the membrane three times 30 min with 1x PBS on a roller.</li>
		<li>Dry the membrane and keep the membrane protected from light using aluminum foil until detection. Membranes can be kept at 4 &#176;C for long term storage.</li>
	</ol>
	</li>
	<li>Image acquisition
	<ol>
		<li>Login to the computer attached to the scanner. Place the membrane on the scanner with the protein side facing down and select the scanning area in the software.</li>
		<li>Optimize the laser intensity for both (700 nm and 800 nm) channels, by confirming no saturation occurs. Acquire the images in both channels and export the images for further analysis (step 6).</li>
	</ol>
	</li>
</ol>
6. Western blot analysis and quantification
NOTE: These recommendations are based on the freely available Image Studio software. However, comparable analyses can also be done using other software packages, such as ImageJ.
<ol>
	<li>Import the file: Create a local workspace on the computer used for analysis. This generates a database of image files for acquired western blots. Import files obtained on the scanner and select the image for analysis.</li>
	<li>TPS analysis
	<ol>
		<li>Display the 700 nm channel to show the total protein staining result. An example of a TPS image is included in Figure 1 in which different tissues from neonatal (P5) (Figure 1A-B) and 10-week old mice (Figure 1C-D) are compared directly. Similarly, Figure 2 shows an example of a direct comparison of brain tissue from mice of different ages.</li>
		<li>Select Analysis tab from top right corner and select Add Rectangle to define the area of interest for normalization (Figure 1B, D). Copy and paste the first rectangle area onto each individual sample to ensure the defined region is at the same size for all analyzed lanes.</li>
		<li>Copy the result from the Shapes tab in the bottom left corner of the software.</li>
	</ol>
	</li>
	<li>Quantification 
	NOTE: The optimal approach for quantifying samples depends on the experimental design. We will here provide an illustrative example of the detection of the survival motor neuron protein (Smn; a key protein involved in the neuromuscular disease spinal muscular atrophy20,21), that is known to decrease over time19 and how normalization of Smn signal intensity to TPS provides reliable estimates of protein expression development.
	<ol>
		<li>Figure 2 shows a decrease of Smn expression with increasing age of the animal with TPS as protein loading control. Repeat Step 6.2.1-6.2.3 to quantify protein loading (Figure 2B). Repeat Step 6.2.1-6.2.3 in the 800 nm channel (Figure 2A) to analyze the protein of interest.</li>
		<li>Copy the results from both TPS and protein of interest to a spreadsheet program. On the spreadsheet, first normalize the protein loading by determining the highest TPS signal and dividing each TPS signal value by this value to obtain the normalized protein loading value.</li>
		<li>Divide the 800 nm signal value from each individual sample by its corresponding normalized protein value to calculate the relative protein expression ratio in different samples.</li>
		<li>After the first normalization, compare various time points or tissues to the average value of the internal standard to allow direct comparisons across different membranes and experiments.</li>
	</ol>
	</li>
</ol>
7. Statistics
<ol>
	<li>To determine any statistically significant differences in protein expression across complex and large groups of samples, appropriate statistical methodology is required. Although a detailed discussion of statistical background goes beyond the scope of this paper, we want to highlight several considerations and detail a successful approach that we have used previously19.</li>
	<li>As for many experiments, protein quantification measurements do not represent completely independent data. Here, for example, multiple tissues are generally obtained from single animals to determine protein levels across multiple organs at a single experimental time-point. Therefore, we used a mixed effects models to analyze differences in protein expression over time, and between tissues.&#160; In general, mixed effects models provide an effective means to deal with non-independence and thereby avoid pseudo-replication22,23. In the present case, mixed effects models increase statistical power by accounting for repeated measurements among tissues, within individuals. We use the statistical software package R to perform these analyses, as this is freely available and versatile. However, other commercially available packages may be able to perform similar analyses. &#160;</li>
	<li>The current experimental design involves a &#8220;split plot&#8221; design because each mouse belonged to only one age group. Therefore, we modeled individual mice (mouse ID) as a random effect with a unique identifier; we also accounted for tissue, age, and their interaction as fixed effects. We modeled the data using the function lmer in the R library, lme4. As a quality control step, we visualized residuals to assess the assumptions of equal variance and a normal distribution and transformed the data where necessary to meet these assumptions. To test for a significant interaction between tissue and age, we fit an identical model that lacked this interaction, and compared the models using parametric bootstrapping (R function PBmodcomp in the library, pbkrtest).&#160; Where significant interactions arose, we used the function emmeans (in the emmeans R library) to determine the cause of the interaction.&#160; For example, we compared mean expression among age classes within each tissue; a significant interaction may arise between age and tissue if, say, two given age classes differed significantly for one tissue but not another. In summary, these approaches provide a way to extend the robustness of conclusions from western blot experiments by providing statistical support to these results.</li>
</ol>

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The authors have no competing interests to disclose.

With the appropriate experimental design, control measures and statistical analysis, western blotting can be used to make reliable quantitative estimates of protein expression within and between a varied range of biological samples. The protocol we describe in the current manuscript aims to serve as a guideline for researchers looking to use Western blotting to undertake quantitative analysis across larger and more complex groups of samples, by using a combination of fluorescence-based detection methods, total protein lo...

Western blotting is a technique that is commonly used to detect and quantify protein expression, including in tissue homogenates or extracts. Over the years, this technique has led to many advances in both basic and clinical research, where it can be used as a diagnostic tool to identify the presence of disease1,2. Western blotting was first described in 1979 as a method to transfer proteins from polyacrylamide gels to nitrocellulose sheets and subsequently visualize proteins using secondary antibodies that were either radioactively labelled or conjugated to fluorescein or peroxidase3. Through the development of commercially available kits and equipment, Western blotting methods have been increasingly standardized and simplified over the years. Indeed, the technique is now readily performed by scientists with varying backgrounds and levels of experience. 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. It is important, therefore, that the accessibility of standardized Western blotting methods does not obscure the need for careful experimental planning and design. Experimental considerations include, but are not limited to, sample preparation and handling, selection and validation of antibodies for protein detection, and gel-to-membrane transfer efficiency of particularly small or large (&#60;10 or &#62;140 kDa) proteins4,5,6,7,8,9. Protein quality of the original sample plays a significant role in determining the outcome of the subsequent Western blot analysis. As protein can be extracted from a wide variety of samples and sources, including cell lines, tissues from animal models, and post-mortem human tissues, consistency in handling and processing is required to obtain reproducible results. For example, when long-term storage of samples for protein extraction is required, it is important to realize that, although protein is generally stable at -80 &#176;C, differences in protein stability between extracted proteins and intact tissues at -80 &#176;C have been reported10. Moreover, to obtain reproducible estimates of protein quantities, consistent homogenization of samples is crucial. Optimizing different lysis buffers and homogenization methods (e.g., manual homogenization compared to automated methods) may be required before starting a large-scale quantitative experiment.
Normalization strategies to correct for protein loading and quantification variability are essential to obtain robust, quantitative results of protein expression. Housekeeping proteins such as &#946;-actin, &#945;-tubulin, &#946;-tubulin, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) have traditionally been used to normalize protein levels for quantification. However, normalization to specific housekeeping proteins for quantification purposes has been increasingly criticized over the past few years11,12. For example, the expression of housekeeping proteins can change across different developmental stages13,14, across tissues from the same animal4, and under various disease conditions4,15,16,17. Therefore, the use of specific housekeeping proteins limits the possibilities of making more complex comparisons between protein expression from different tissues, at different timepoints and under varying experimental conditions. An alternative to housekeeping proteins to control for protein loading variation is the use of a total protein stain (TPS) that labels and visualizes all proteins present in a sample. TPS allows signal normalization based on total protein load rather than levels of one specific protein and therefore quantification of TPS signal should be comparable and reproducible regardless of experimental condition, sample type or developmental timepoint. Examples of total protein stains include Ponceau S, stain-free gels, Coomassie R-350, Sypro-Ruby, Epicocconone, Amydo Black, and Cy5 (reviewed in ref. 18). Each of these methods has specific advantages and limitations and method selection depends on the time and tools available as well as the experimental setup4,18.
In addition to using a TPS to correct for within-membrane loading and quantification variability, it may be necessary to compare samples between different membranes, particularly when performing large-scale protein expression analysis. However, variability in factors such as antibody binding efficiency and total protein stain intensity may introduce further variability between protein samples that are analyzed on separate gels and membranes. For robust quantification in this situation, it is therefore necessary to introduce a further normalization step to account for between-membrane variability. This can be achieved by including an internal loading standard on each of the separately analyzed membranes that is kept constant across experiments. This standard can take the form of any protein lysate that can be obtained in sufficient quantities to be used across all membranes included in the experiment. Here, we use a lysate of mouse brain (obtained from 5 day old control mice), as brain is readily homogenized and the obtained protein lysate contains a significant amount of protein at a high concentration. Loading an internal standard in triplicate allows samples on separate membranes to be normalized and compared directly.
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 timepoints19. By combining a fluorescent TPS with the use of an internal loading standard, we were able to overcome existing limitations in the number of samples and experimental conditions that can be compared within a single experiment. This approach expands the use of traditional Western blot techniques, thereby allowing researchers to better explore protein expression across different tissues and samples.

<table><tbody><tr><td>Fine Tipped Gel Loading Tips</td><td>Alpha Laboratories</td><td>GL20057SNTL</td><td></td></tr><tr><td>Halt Protease Inhibitor Cocktail, EDTA-free 100x 5mL</td><td>ThermoFisher Scientific</td><td>78437</td><td></td></tr><tr><td>Handheld homogeniser&amp;nbsp;</td><td>VWR Collection</td><td>431-0100</td><td></td></tr><tr><td>iBlot 2 Gel Transfer Device</td><td>ThermoFisher Scientific</td><td>IB21001</td><td></td></tr><tr><td>iBlot Transfer Stack, PVDF, regular size</td><td>ThermoFisher Scientific</td><td>IB401031</td><td></td></tr><tr><td>Image Studio Lite</td><td>Licor</td><td>N/A</td><td>Free download from https://www.licor.com/bio/products/software/image_studio_lite/</td></tr><tr><td>IRDye 800CW secondary antibodies</td><td>Licor</td><td>--</td><td>Select appropriate secondary antibody that is specific against host of primary antibody.</td></tr><tr><td>Micro BCA Protein Assay Kit</td><td>ThermoFisher Scientific</td><td>23235</td><td></td></tr><tr><td>Novex Sharp Pre-stained Protein Standard</td><td>ThermoFisher Scientific</td><td>LC5800</td><td></td></tr><tr><td>NuPAGE 4-12% Bis-Tris Protein Gels, 1.0 mm, 15-well</td><td>ThermoFisher Scientific</td><td>NP0323BOX</td><td></td></tr><tr><td>NuPAGE LDS Sample Buffer (4X)</td><td>ThermoFisher Scientific</td><td>NP0007</td><td></td></tr><tr><td>NuPAGE MOPS SDS Running Buffer (20X)</td><td>ThermoFisher Scientific</td><td>NP0001</td><td></td></tr><tr><td>Odyssey Blocking Buffer</td><td>Licor</td><td>927-40000</td><td></td></tr><tr><td>Purified Mouse anti-SMN (survival motor neuron) monoclonal antibody</td><td>BD Transduction Laboratories</td><td>610646</td><td>Is used extensively in the SMN/SMA literature and gives consistent results regardsless of lot number</td></tr><tr><td>REVERT Total Protein Stain, 250 mL&amp;nbsp;</td><td>Licor</td><td>926-11021&amp;nbsp;</td><td></td></tr><tr><td>REVERT Wash Solution&amp;nbsp;</td><td>Licor</td><td>926-11012&amp;nbsp;</td><td></td></tr><tr><td>RIPA Lysis and Extraction Buffer</td><td>ThermoFisher Scientific</td><td>89900</td><td></td></tr><tr><td>XCell&amp;nbsp;SureLock Mini-Cell</td><td>ThermoFisher Scientific</td><td>EI0001</td><td></td></tr></tbody></table>

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.

We include examples of the use of TPS and an internal standard to facilitate comparisons of protein levels across tissues and time points. Figure 1 shows results from Western blotting on protein extracted from tissues obtained from neonatal (postnatal day 5) in comparison to adult mice (10-week old). TPS and Smn immunoblot are shown in Figure 1A, C. Quantification of fluorescence intensity of the TPS was achieved...

Watch this Scientific Journal Video about Robust Comparison of Protein Levels Across Tissues and Throughout Development Using Standardized Quantitative Western Blotting at JoVE.com

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

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 ...

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Biochemistry

14.1K Views.  University of Edinburgh. 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.

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

High Throughput, Absolute Determination of the Content of a Selected Protein at Tissue Levels Using Quantitative Dot Blot Analysis (QDB)

Lacking a convenient, quantitative, high throughput immunoblot method for absolute determination of the content of a specific protein at cellular and tissue level significantly hampers the progress in proteomic research. Results derived from currently available immunoblot techniques are also relative, preventing any efforts to combine independent studies with a large-scale analysis of protein samples. In this study, we demonstrate the process of quantitative dot blot analysis (QDB) to achieve absolute quantification in a high throughput format. Using a commercially available protein standard, we are able to determine the absolute content of capping actin protein, gelsolin-like (CAPG) in protein samples prepared from three different mouse tissues (kidney, spleen, and prostate) together with a detailed explanation of the experimental details. We propose the QDB analysis as a convenient, quantitative, high throughput immunoblot method of absolute quantification of individual proteins at the cellular and tissue level. This method will substantially aid biomarker validation and pathway verification in various areas of biological and biomedical research.

High Throughput, Absolute Determination of the Content of a Selected Protein at Tissue Levels Using Quantitative Dot Blot Analysis QDB

Here we demonstrate a detailed process of quantitative dot blot analysis (QDB) by determining the absolute content of a targeted protein, capping actin protein, gelsolin-like (CAPG), in three different mouse tissues. We demonstrate a high throughput, convenient, quantitative immunoblot technique for biomarker validation at the cellular and tissue level.

Lacking a convenient, quantitative, high throughput immunoblot method for absolute determination of the content of a specific protein at cellular and ...

Consensus Brain-derived Protein, Extraction Protocol for the Study of Human and Murine Brain Proteome Using Both 2D-DIGE and Mini 2DE Immunoblotting

Two-dimensional gel electrophoresis (2DE) is a powerful tool to uncover proteome modifications potentially related to different physiological or pathological conditions. Basically, this technique is based on the separation of proteins according to their isoelectric point in a first step, and secondly according to their molecular weights by SDS polyacrylamide gel electrophoresis (SDS-PAGE). In this report an optimized sample preparation protocol for little amount of human post-mortem and mouse brain tissue is described. This method enables to perform both two-dimensional fluorescence difference gel electrophoresis (2D-DIGE) and mini 2DE immunoblotting. The combination of these approaches allows one to not only find new proteins and/or protein modifications in their expression thanks to its compatibility with mass spectrometry detection, but also a new insight into markers validation. Thus, mini-2DE coupled to western blotting permits to identify and validate post-translational modifications, proteins catabolism and provides a qualitative comparison among different conditions and/or treatments. Herein, we provide a method to study components of protein aggregates found in AD and Lewy body dementia such as the amyloid-beta peptide and the alpha-synuclein. Our method can thus be adapted for the analysis of the proteome and insoluble proteins extract from human brain tissue and mice models too. In parallel, it may provide useful information for the study of molecular and cellular pathways involved in neurodegenerative diseases as well as potential novel biomarkers and therapeutic targets.

A common protein extraction protocol using urea/thiourea/SDS buffer for human and mice brain tissue allows indentification of proteins by 2D-DIGE and their subsequent characterization by mini 2DE immunoblotting. This method enables one to obtain more reproducible and reliable results from human biopsies and experimental models.

Two-dimensional gel electrophoresis (2DE) is a powerful tool to uncover proteome modifications potentially related to different physiological or ...

Neuroscience

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.

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 ...

Biology

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 ...

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to facilitate sensory functions in cells. The level of a protein may be regulated differently at centrosomes than at other .cellular locations, and the variation in the centrosomal level of several proteins at different points of the cell cycle appears to be crucial for the proper regulation of centriole assembly. We developed a quantitative fluorescence microscopy assay that measures relative changes in the level of a protein at centrosomes in fixed cells from different samples, such as at different phases of the cell cycle or after treatment with various reagents. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region, and normalize that measurement against the same for another protein that does not vary under the chosen experimental condition. Utilizing this assay in combination with BrdU pulse and chase strategy to study unperturbed cell cycles, we have quantitatively validated our recent observation that the centrosomal pool of VDAC3 is regulated at centrosomes during the cell cycle, likely by proteasome-mediated degradation specifically at centrosomes.

Here, a novel quantitative fluorescence assay is developed to measure changes in the level of a protein specifically at centrosomes by normalizing that protein&#8217;s fluorescence intensity to that of an appropriate internal standard.

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to ...

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 ...

Quantitative Immunoblotting of Cell Lines as a Standard to Validate Immunofluorescence for Quantifying Biomarker Proteins in Routine Tissue Samples

Quantification of proteins of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples is important in clinical and research applications. An optimal method of quantification is accurate, has a broad linear dynamic range and maintains the structural integrity of the sample to allow for identification of individual cell types. Current methods such as immunohistochemistry (IHC), mass spectrometry, and immunoblotting each fail to meet these stipulations due to their categorical nature or need to homogenize the sample. As an alternative method, we propose the use of immunofluorescence (IF) and image analysis to determine the relative abundance of a protein of interest in FFPE tissues. Herein we demonstrate that this method is easily optimized, yields a wide dynamic range, and is linearly quantifiable as compared to the gold standard of quantitative immunoblotting. Furthermore, this method permits the maintenance of the structural integrity of the sample and allows for the distinction of various cell types, which may be crucial in diagnostic applications. Overall, this is a robust method for the relative quantification of proteins in FFPE samples and can be easily adapted to suit clinical or research needs.

We describe the use of quantitative immunoblotting to validate immunofluorescence histology coupled with image analysis as a means of quantifying a protein of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples. Our results demonstrate the utility of immunofluorescence histology for ascertaining the relative quantity of biomarker proteins in routine biopsy samples.

Quantification of proteins of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples is important in clinical and research applications. An ...

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 ...

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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.

Source: Huang, Y. T., et al. Robust Comparison of Protein Levels Across Tissues and Throughout Development Using Standardized Quantitative Western ...

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Biological Techniques

Blot Techniques

Western Blotting

Western blotting is an analytical technique for protein identification. It has various applications in immunology and medicine, including detecting diseases like bovine spongiform encephalopathy, mad cow disease, and human and feline immunodeficiency virus from biological samples.
The technique begins with separating proteins from the sample using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), followed by protein transfer, immunoblotting, and finally, protein detection.
The proteins from the polyacrylamide gel are transferred to specialized membranes like nitrocellulose (NC) or polyvinylidene difluoride (PVDF). Compared to NC, PVDF has higher durability and protein binding capacity and can be re-probed.
The protein transfer is either by electric current application (electroblotting) or capillary action. In electroblotting, the hydrophobic interactions transfer the negatively charged proteins from the gel and trap them on the membrane. In the other method, a filter paper placed on the gel soaks the buffer by capillary action, simultaneously drawing the proteins from the gel and transferring them to the membrane. To ensure the quality of transferred bands, non-specific protein stains like Ponceau S allow for a reversible check.
The next step, immunoblotting, involves treating the membrane with a blocking buffer like 3-5% Bovine serum albumin or non-fat dried milk. These buffers reduce the chance of non-specific binding of the primary antibody to the membrane. The primary antibodies can either be polyclonal or monoclonal and bind to the protein bands on the membrane through their Fab region. Next, the added secondary antibodies bind and attach to the Fc region of the primary antibodies. The secondary antibodies are tagged with enzymes which, upon suitable substrate addition, produce detectable colorimetric or chemiluminescent signals. Further, densitometry techniques can quantify the optical density of these imaged bands, corresponding to the abundance of proteins in the sample.
The expected result for a western blot can be affected by various factors like erroneous sample loading, inaccurate transfer with trapped air bubbles, and the use of non-specific antibodies. Also, inappropriate primary and secondary antibody concentrations and contaminated buffers can affect the quality of bands and their visualization.

Western blotting is an analytical technique for protein identification. It has various applications in immunology and medicine, including detecting ...