protein blotting

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Western blot From Wikipedia, the free encyclopedia Western blot using an antibody that recognizes proteins modified with lipoic acid. The western blot (sometimes called the protein immunoblot) is a widely used analytical technique used to detect specific proteins in a sample of tissue homogenate or extract. It uses gel electrophoresis to separate native proteins by 3-D structure or denatured proteins by the length of the polypeptide. The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are stained with antibodies specific to the target protein. [1][2] The gel electrophoresis step is included in Western blot analysis to resolve the issue of the cross-reactivity of antibodies. An improved immunoblot method, Zestern analysis, is able to address this issue without the electrophoresis step, thus significantly improving the efficiency of protein analysis. There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against tens of thousands of different proteins. [3] Commercial antibodies can be expensive, although the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines. Other related techniques include dot blot analysis, zestern analysis, immunohistochemistry where antibodies are used to detect proteins in tissues and cells by immunostaining, and enzyme-linked immunosorbent assay (ELISA). Steps[edit ] Tissue preparation[edit ] Samples can be taken from whole tissue or from cell culture. Solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonication . Cells may also be broken open by one of the above

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Page 1: Protein Blotting

Western blotFrom Wikipedia, the free encyclopedia

Western blot using an antibody that recognizes proteins modified with lipoic acid.

The western blot (sometimes called the protein immunoblot) is a widely used analytical technique used to detect specific proteins in a sample of tissue homogenate or extract. It uses gel electrophoresis to separate native proteins by 3-D structure or denatured proteins by the length of the polypeptide. The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are stained with antibodies specific to the target protein.[1][2] The gel electrophoresis step is included in Western blot analysis to resolve the issue of the cross-reactivity of antibodies. An improved immunoblot method, Zestern analysis, is able to address this issue without the electrophoresis step, thus significantly improving the efficiency of protein analysis.

There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against tens of thousands of different proteins.[3] Commercial antibodies can be expensive, although the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines.

Other related techniques include dot blot analysis, zestern analysis, immunohistochemistry where antibodies are used to detect proteins in tissues and cells by immunostaining, and enzyme-linked immunosorbent assay (ELISA).

Steps[edit]

Tissue preparation[edit]

Samples can be taken from whole tissue or from cell culture. Solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer(smaller volumes), or by sonication. Cells may also be broken open by one of the above mechanical methods. However, virus or environmental samples can be the source of protein and thus western blotting is not restricted to cellular studies only.

Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. Protease and phosphatase inhibitors are often added to prevent the digestion of the sample by its own enzymes. Tissue preparation is often done at cold temperatures to avoid protein denaturing and degradation.

A combination of biochemical and mechanical techniques – comprising various types of filtration and centrifugation – can be used to separate different cell compartments andorganelles.

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Gel electrophoresis[edit]

Main article: Gel electrophoresis

The proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. The nature of the separation depends on the treatment of the sample and the nature of the gel. This is a very useful way to identify a protein.

By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers loaded with sodium dodecyl sulfate (SDS). SDS-PAGE (SDS polyacrylamide gel electrophoresis) maintains polypeptides in a denatured state once they have been treated with strong reducing agents to remove secondary and tertiary structure (e.g. disulfide bonds [S-S] to sulfhydryl groups [SH and SH]) and thus allows separation of proteins by their molecular weight. Sampled proteins become covered in the negatively charged SDS and move to the positively charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster through this mesh and the proteins are thus separated according to size (usually measured in kilodaltons, kDa). The concentration of acrylamide determines the resolution of the gel - the greater the acrylamide concentration the better the resolution of lower molecular weight proteins. The lower the acrylamide concentration the better the resolution of higher molecular weight proteins. Proteins travel only in one dimension along the gel for most blots.

Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder, a commercially available mixture of proteins having defined molecular weights, typically stained so as to form visible, coloured bands. When voltage is applied along the gel, proteins migrate through it at different speeds dependent on their size. These different rates of advancement (different electrophoretic mobilities) separate into bands within each lane.

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It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single sample out in two dimensions. Proteins are separated according to isoelectric point (pH at which they have neutral net charge) in the first dimension, and according to their molecular weight in the second dimension.

Transfer[edit]

In order to make the proteins accessible to antibody detection they are moved from within the gel onto a membrane made of nitrocellulose or polyvinylidene difluoride (PVDF). The primary method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The proteins move from within the gel onto the membrane while maintaining the organization they had within the gel. An older method of transfer involves placing a membrane on top of the gel, and a stack of filter papers on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. In practice this method is not used as it takes too much time; electroblotting is preferred. As a result of either "blotting" process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.

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The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie Brilliant Blue or Ponceau Sdyes. Ponceau S is the more common of the two, due to its higher sensitivity and water solubility, the latter making it easier to subsequently destain and probe the membrane, as described below.[6]

Blocking[edit]

Since the membrane has been chosen for its ability to bind protein and as both antibodies and the target are proteins, steps must be taken to prevent the interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically 3-5% Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive) in Tris-Buffered Saline (TBS) or I-Block, with a minute percentage (0.1%) of detergent such as Tween 20 or Triton X-100. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the western blot, leading to clearer results, and eliminates false positives.

Detection[edit]

During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme; when exposed to an appropriate substrate this enzyme drives a colourimetric reaction and produces a color. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.

Two steps[edit]

Primary antibody

The primary antibodies are generated when a host species or immune cell culture is exposed to protein of interest (or a part thereof). Normally, this is part of the immune response, whereas

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here they are harvested and used as sensitive and specific detection tools that bind the protein directly.

After blocking, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/mL) is incubated with the membrane under gentle agitation. Typically, the solution is composed of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with higher temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").

Secondary antibody

After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. Antibodies come from animal sources (or animal sourced hybridoma cultures); an anti-mouse secondary will bind to almost any mouse-sourced primary antibody, which allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. This is known as a secondary antibody, and due to its targeting properties, tends to be referred to as "anti-mouse," "anti-goat," etc. The secondary antibody is usually linked to biotin or to a reporter enzyme such asalkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhance the signal.

Most commonly, a horseradish peroxidase-linked secondary is used to cleave a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot. A cheaper but less sensitive approach utilizes a 4-chloronaphthol stain with 1% hydrogen peroxide; reaction of peroxide radicals with 4-chloronaphthol produces a dark purple stain that can be photographed without using specialized photographic film.

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As with the ELISPOT and ELISA procedures, the enzyme can be provided with a substrate molecule that will be converted by the enzyme to a colored reaction product that will be visible on the membrane (see the figure below with blue bands).

Another method of secondary antibody detection utilizes a near-infrared (NIR) fluorophore-linked antibody. Light produced from the excitation of a fluorescent dye is static, making fluorescent detection a more precise and accurate measure of the difference in signal produced by labeled antibodies bound to proteins on a western blot. Proteins can be accurately quantified because the signal generated by the different amounts of proteins on the membranes is measured in a static state, as compared to chemiluminescence, in which light is measured in a dynamic state. [7]

A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A or Streptavidin with a radioactive isotope of iodine. Since other methods are safer, quicker, and cheaper, this method is now rarely used; however, an advantage of this approach is the sensitivity of auto-radiography based imaging, which enables highly accurate protein quantification when combined with optical software (e.g. Optiquant).

One step[edit]

Historically, the probing process was performed in two steps because of the relative ease of producing primary and secondary antibodies in separate processes. This gives researchers and corporations huge advantages in terms of flexibility, and adds an amplification step to the detection process. Given the advent of high-throughput protein analysis and lower limits of detection, however, there has been interest in developing one-step probing systems that would allow the process to occur faster and with fewer consumables. This requires a probe antibody which both recognizes the protein of interest and contains a detectable label, probes which are often available for known protein tags. The primary probe is incubated with the membrane in a manner similar to that for the primary antibody in a two-step process, and then is ready for direct detection after a series of wash steps.

Western blot using radioactive detection system

Analysis[edit]

After the unbound probes are washed away, the western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is repeated for a structural protein, such as actin or tubulin, that should not change between

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samples. The amount of target protein is normalized to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.

Colorimetric detection[edit]

The colorimetric detection method depends on incubation of the western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) orspectrophotometry.

Chemiluminescent detection[edit]

Chemiluminescent detection methods depend on incubation of the western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by photographic film, and more recently by CCD cameras which capture a digital image of the western blot. The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used.

Radioactive detection[edit]

Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is declining due to its hazardous radiation[citation

needed], because it is very expensive, health and safety risks are high, and ECL (enhanced chemiluminescence) provides a useful alternative.

Fluorescent detection[edit]

The fluorescently labeled probe is excited by light and the emission of the excitation is then detected by a photosensor such as CCD camera equipped with appropriate emission filters

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which captures a digital image of the western blot and allows further data analysis such as molecular weight analysis and a quantitative western blot analysis. Fluorescence is considered to be one of the best methods for quantification, but is less sensitive than chemiluminescence.[8]

Secondary probing[edit]

One major difference between nitrocellulose and PVDF membranes relates to the ability of each to support "stripping" antibodies off and reusing the membrane for subsequent antibody probes. While there are well-established protocols available for stripping nitrocellulose membranes, the sturdier PVDF allows for easier stripping, and for more reuse before background noise limits experiments. Another difference is that, unlike nitrocellulose, PVDF must be soaked in 95% ethanol, isopropanol or methanol before use. PVDF membranes also tend to be thicker and more resistant to damage during use.

2-D gel electrophoresis[edit]

Main article: Two-dimensional gel electrophoresis

2-dimensional SDS-PAGE uses the principles and techniques outlined above. 2-D SDS-PAGE, as the name suggests, involves the migration of polypeptides in 2 dimensions. For example, in the first dimension polypeptides are separated according to isoelectric point, while in the second dimension polypeptides are separated according to their molecular weight. The isoelectric point of a given protein is determined by the relative number of positively (e.g. lysine and arginine) and negatively (e.g. glutamate and aspartate) charged amino acids, with negatively charged amino acids contributing to a low isoelectric point and positively charged amino acids contributing to a high isoelectric point. Samples could also be separated first under nonreducing conditions using SDS-PAGE and under reducing conditions in the second dimension, which breaks apart disulfide bonds that hold subunits together. SDS-PAGE might also be coupled with urea-PAGE for a 2-dimensional gel.

In principle, this method allows for the separation of all cellular proteins on a single large gel. A major advantage of this method is that it often distinguishes between differentisoforms of a particular protein - e.g. a protein that has been phosphorylated (by addition of a negatively charged group). Proteins that have been separated can be cut out of the gel and then analysed by mass spectrometry, which identifies the protein.

Please refer to reference articles for examples of the application of 2-D SDS PAGE.

Medical diagnostic applications[edit]

The confirmatory HIV test employs a western blot to detect anti-HIV antibody in a human serum sample. Proteins from known HIV-infected cells are separated and blotted on a membrane as above. Then, the serum to be tested is applied in the primary antibody incubation step; free antibody is washed away, and a secondary anti-human antibody linked to an enzyme signal is added. The stained bands then indicate the proteins to which the patient's serum contains antibody.

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A western blot is also used as the definitive test for Bovine spongiform encephalopathy (BSE, commonly referred to as 'mad cow disease').

Some forms of Lyme disease testing employ western blotting.

Western blot can also be used as a confirmatory test for Hepatitis B infection.

In veterinary medicine, western blot is sometimes used to confirm FIV+ status in cats

Western blotting (immunoblot): Gel electrophoresis for proteinsWestern Blotting (also called immunoblotting) is a technique used for analysis of individual proteins in a protein mixture (e.g. a cell lysate). In Western blotting (immunoblotting) the protein mixture is applied to a gel electrophoresis in a carrier matrix (SDS-PAGE, native PAGE, isoelectric focusing, 2D gel electrophoresis, etc.) to sort the proteins by size, charge, or other differences in individual protein bands. The separated protein bands are then transferred to a carrier membrane (e.g. nitrocellulose, nylon or PVDF). This process is called blotting. The proteins adhere to the membrane in the same pattern as they have been separated due to interactions of charges. The proteins on this immunoblot are then accessible for antibody binding for detection.

 An

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tibodies are used to detect target proteins on the western blot (immunoblot). The antibodies are conjugated with fluorescent or radioactive labels or enzymes that give a subsequent reaction with an applied reagent, leading to a coloring or emission of light, enabling detection.

The term Western blotting is based on a play of words. The southern blot, which is a method to detect specific DNA sequences, is named after Ed Southern, who first described this procedure. The western blot (immunoblot), as well as the northern blot (for RNA detection), play on the meaning of this name.

What type of gel electophoresis for proteins are possible?

One can choose from different types of gel electrophoresis for proteins depending on the criteria by which the proteins should be separated. Some commonly used electrophoretic methods are: SDS-PAGE, native-PAGE and isoelectric focusing.

SDS-PAGE: This is a denaturing method as it treats the proteins with anionic SDS detergent (sodiumdodcylsulfate). Secondary- and tertiary structure are destroyed by this process. Additionally, SDS binds the proteins and thereby covers their chemical charges, leading to equally negatively charged proteins. Therefore the following separation happens solely by the size of the polypeptide chains in the polyacrylamide gel.

Native PAGE: Native, unfolded, and not-denatured proteins can be separated using this method. This method allows for the separation of proteins that are inaccessible by other methods. One example would be the separation of modified and unmodified proteins of the same kind (e.g. phosphorylated versus unphosphorylated state of a protein). Native PAGE can also be used to confirm biologically relevant conformations, like di-, tri-, or tetrameric forms of proteins (contrary to SDS-PAGE, which would separate the individual and denatured peptide chains). This method can also detect different complexes of different proteins.

The separation using native PAGE depends on a number of parameters such as the charge, size and 3D structure of the protein. A suitable buffer is needed to maintain the 3D folding of the protein. The applicability of the buffer depends on the isoelectric point and the charges of the protein.

Isoelectric focusing: This method builds on the fact that a protein has a specific charge at certain pH values. Depending on the pH the acidic and basic functional groups contribute by increasing or decreasing the total charge of the protein. The isoelectric point is defined as the the point where the total charge of the molecule is zero, because there is an equal amount of negative and positive charges in the molecule.

Special gradient gels are needed for isoelectric focusing as the pH changes from acidic to basic along a gradient within the gel. Due to an electric charge connected to the gel the protein travels to the point in the gel where the charge of the gel equals that of the protein, and the total charge equals zero, i.e. the isoelectric point. Hence, this method is used to separate proteins by their charges, as well as to determine the isoelectric point of a target protein. The separation occurs due to the charge of the protein or by the number of basic- and acidic groups the protein contains.

The above-mentioned methods for gel electrophoresis of proteins can also be combined to separate proteins. The choice of methods depends on the specific requirements of the experiment.

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Blotting

Following the separation of the protein mix the polypeptide bands are transferred to a membrane carrier. For this purpose the membrane is attached to the gel and this so-called sandwich is transferred to an electrophoresis chamber. It is possible that some of the SDS is washed out, and the protein partially re-naturates again, i.e. regains its 2D- and 3D structure. However, the applied electric charge causes the proteins to travel out of the gel vertically to the direction they traveled in on the gel, onto the membrane. The protein bands are thereby bound to the membrane. The "blotted" bands are now available to be treated further (e.g. for detection of specific proteins with specific antibodies).

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Immunodetection

The identification of specific antibodies is possible after the separation and blotting of proteins. Specific antibodies (mono- or polyclonal) bind to "their" band of proteins. Unspecifically binding antibodies are removed by washing with detergent-containing buffers. Additionally, unspecific binding pockets can be blocked before the addition of specific antibodies.

Primary antibodies are usually applied first, which are then recognized by a secondary antibody. The secondary antibody is conjugated with colour, radioactivity or an enzyme for detection. Biotin-conjugated antibodies are also used for this purpose.

It can occasionally be advantageous to use polyclonal primary antibodies as such antibodies recognize several epitopes, contrary to monoclonal antibodies that are restricted in their binding affinity. After immunodetection it is possible to strip the antibody off the membrane for further analysis with other antibodies (e.g. in order to detect other specific antibodies from the protein mixture under investigation).

Analysis of the western blot is then carried out using a variety of different imaging systems (e.g. luminescence, color reaction, autoradiography).

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Why western blotting (immunoblot)?

The western blotting (immunoblot) method entails various advantages as compared to other immunosorbent assays (ISAs), like for example ELISA.

Western blotting (immunoblot) expands on the idea of ELISA by allowing separation of the protein mix by size, charge, and/or conformation. The described method of stripping allows for the detection of several targets, contrary to ELISA where only one protein can be detected. As the gel electophoreis of proteins separates the proteins into bands, one can determine the size of the target protein/polypeptide. It is also possible to (semi-)quantify the protein of interest by running an internal quantity standard in parallel with the samples in the gel. Similarly, the protein content of the samples can be compared ("sample A contains more protein than sample B").

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A disadvantage of western blotting (immunoblot) is that it is time-consuming (compared to ELISA) and has a high demand in terms of experience of the experimenter. Additionally, it requires optimizing the experimental conditions (i.e. protein isolation, buffers, type of separation, gel concentration, etc.).

There are many different types and methods for western blotting (immunoblot). Hence, it covers very different topics and applications.

Hint: Browse through almost 260,000 antibodies for western blotting (immunoblot)

12/06/2013 | David Kitz Kramer (updated by Eri Kinoshita) Applications

General Monoclonal antibodies Secondary antibodies anti-Immunoglobulin G (IgG) secondary antibodies anti-Immunoglobulin A (IgA) secondary antibodies anti-Immunoglobulin E (IgE) secondary antibodies anti-Immunoglobulin D (IgD) secondary antibodies anti-Immunoglobulin M (IgM) secondary antibodies anti-Immunoglobulin Y (IgY) secondary antibodies

Applications Radioimmunoassay (RIA) Immunohistochemistry (IHC) Loading controls Isotype control antibodies Western blotting (immunoblot): Gel electrophoresis for proteins Blocking peptides Tissue lysates Whole cell lysates Enzyme-linked immunosorbent assay (ELISA) What is flow cytometry (FACS analysis)? General flow cytometry protocol Antibody purification using antibody-binding proteins ChromoTek Chromobodies®: Novel species of intracellular functional antibodies Metabolism assay kits from Biovision available on antibodies-online.com GFP-Trap® GFP-Booster Antibody labeling kits Shenandoah's stem cell growth factors

Research Areas Cytoskeleton Proteases

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Autophagy Type 2 Diabetes mellitus (T2D) Metabolism JNK pathway Cytokeratins in the detection of tumors Apoptosis Extracellular matrix Major Histocompatability Complex (MHC) Leukemia Heat-Shock Proteins

Protein blot (Western)

A protein blot, also known as the immunoblot or Western blot, is a method of semiquantitative determination of protein expression. Crude cell lysates are loaded into a polyacrylamide gel containing a denaturing agent which give all the proteins a net negative charge. A current passing through the gel will then propel the proteins through the gel, with the largest proteins travelling the slowest. This results in the proteins being roughly separated by their mass. The proteins are then transferred from the gel onto a membrane (often nitrocellulose or PVDF), which is then incubated with an antibody directed against the protein of interest. By using a detector conjugated to an antibody one can then specifically detect the protein of interest.

Standard western blot procedure

Transient SLUG gene expression in 293GP cells (last lane); titration of the primary and adjusting exposure time will

improve the signal:noise

Prior to running the western blotting experiment, consult with the vendor in order to determine if they have optimized the titration for the antibody you just purchased. Most vendors should offer some

Page 14: Protein Blotting

guideline for how to use the antibody. Value your time and effort first and foremost. If the experiment comes out strange, make the call or email the vendor for assistance. Your time is the most valuable element to the procedure!

Mouse monoclonal western blot

Load at least 50 ug of whole cell lysate or 20 ug nuclear extract prepared from fresh buffers (30 µg of protein ~ 25,000 cells). Resolve proteins on the gel voltage gradient.

Transfer proteins to a nitrocellulose or PVDF membrane and block the membrane in 10 cm square dish with fresh 5% milk TTBS 2 hr room temperature, or overnight at 4C.

Perform 3-5 shake rinses with cold TTBS (until no residual milk is observed). Incubate the primary antibody 1:200 in fresh 3% milk TTBS for 90 minutes room temperature. Perform 3 shake rinses, followed by 4X for 5 minutes washes with ambient or cold TTBS. Incubate the the secondary antibody 1:1000 anti-mouse in fresh 1% milk TTBS for 60 minutes at

room temperature. Perform 3 shake rinses, followed by 4X for 5 minutes washes with ambient or cold TTBS. Perform ECL using a suitable detection reagent. Wick away all residual chemiluminescent liquid

by dabbing membrane edges on paper towel. Prepare the membranes for ECL by covering with either saran wrap, or a transparency page.

There should be no running of ECL liquid. Optimize protein signal with multiple exposure times.Goat polyclonal western blot

Load at least 50 ug of whole cell lysate or 20 ug nuclear extract prepared from fresh buffers (30 µg of protein ~ 25,000 cells). Resolve proteins on the gel voltage gradient.

Transfer proteins to a nitrocellulose or PVDF membrane and block the membrane in 10 cm square dish with fresh 5% milk TTBS 2 hr room temperature, or overnight at 4C.

Perform 3-5 shake rinses (until very little residual milk washes off the membrane). Incubate membrane with the primary antibody 1:200 in fresh 5% Milk TTBS for 90 minutes room

temperature. Perform 3 shake rinses, followed by 4X 5 minute wash with ambient or cold TTBS. Incubate the the secondary antibody 1:5000 anti-goat in fresh 5% Milk TTBS for 60 minutes at

room temperature. Perform shake rinses, until no residual milk is observed, followed by 4X 5 minute wash with

ambient or cold TTBS. Perform ECL using a suitable detection reagent. Wick away all residual chemiluminescent liquid

by dabbing membrane edges on paper towel. Prepare the membranes for ECL by covering with either saran wrap, or a transparency page.

There should be no running of ECL liquid. Optimize protein signal with multiple exposure times.Rabbit polyclonal western blot

Load at least 50 ug of whole cell lysate or 20 ug nuclear extract prepared from fresh buffers (30 µg of protein ~ 25,000 cells). Resolve proteins on the gel voltage gradient.

Transfer proteins to a nitrocellulose or PVDF membrane and block in 10 cm square dish with fresh 5% milk TTBS 2 hr, r.t. or overnight at 4C.

Perform 3 shake rinses (until very little residual milk washes off the membrane).

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Incubate the primary antibody 1:200 in fresh 3% milk TTBS for 90 minutes room temperature. Perform 3 shake rinses, followed by 4X 5 minute wash with ambient or cold TTBS. Incubate the the secondary antibody 1:4000 anti-rabbit in fresh 1% Milk TTBS for 60 minutes at

room temperature. Perform 3 shake rinses, followed by 4X 5 minute wash with ambient or cold TTBS. Perform ECL using a suitable detection reagent. Wick away all residual chemiluminescent liquid

by dabbing membrane edges on paper towel. Prepare the membranes for ECL by covering with either saran wrap, or a transparency page.

There should be no running of liquid. Optimize protein signal with multiple exposure times.Phosphorylation state specific western blotAdjusting certain incubation conditions can improve the detection signal for the phospho-specific antibody. When serine/threonine phosphatase inhibitor Sodium Fluoride (NaF), and tyrosine phosphatase inhibitor Sodium Orthovanadate (Na3VO4) are included in the blocking and incubation buffers, phospho-specific signals can noticeably improve. Including 50 mM NaF and 5 mM Na3VO4 in the blocking and incubation buffers can improve the signal. Using 5% milk diluent for primary and secondary incubations will reduce the nonspecific banding and background.

Load 50 ug of whole cell lysate/tissue extract OR 20 ug of nuclear extract prepared from fresh buffers. Resolve proteins on the gel voltage gradient.

Transfer proteins to a nitrocellulose or PVDF membrane and block in fresh 5% milk TTBS, 50 mM NaF/5 mM Na3VO4 for 2 hr, r.t. or overnight at 4C. Perform 3 shake rinses and drain until no residual milk appears to stream off the wash buffer.

Incubate the primary antibody 1:200-20000 in fresh 5% milk TTBS, 50 mM NaF/5 mM Na3VO4 for 120 minutes room temperature. Perform 3 shake rinses followed by wash with cold TTBS 4X for 5 minutes each wash.

Incubate the the secondary antibody 1:2000 anti-mouse, 1:5000 anti-rabbit, 1:5000 anti-goat in fresh 5% milk TTBS, 50 mM NaF/5 mM Na3VO4 for 60 minutes at room temperature. Perform 3 shake rinses followed by wash with cold TTBS 4X for 5 minutes each wash.

Perform ECL using a suitable chemiluminescent reagent Optimize protein signal with multiple exposure times.Dephosphorylation of Lysate with Lambda PhosphataseLysate Treatment

To a micro-centrifuge tube add 80 µg of lysate. Amount maybe scaled up or down as needed. Add 6 microliters of lambda-PPase Buffer. Final concentration: 50 mM Tris, pH 7.5, 0.1 mM

Na2EGTA, 5 mM dithiothreitol, 2 mM MnCl2. Add 400 units of l-Phosphatase

Santa Cruz Biotechnology Inc. sc-200312

Millipore Cat. #14-405

Incubate at 30˚C for 20 minutes. The protein sample/lysate is now ready for analysis or further preparation depending on

application

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1. Sharma SK and Carew TJ. .  pmid:12137799. PubMed HubMed [Paper1]Membrane Treatment

Treats ~30 lanes or 2-3 (6x12 cm) membranes. Mark treated and untreated membranes.

Block membranes in 3%BSA, 1X TBS, 0.1% Triton X-100. Add 15 ul of Lambda-PPase (Santa Cruz Biotechnology Inc. sc-200312) in 15 ml of blocking

buffer + 2mM DTT, 2mM MnCl2. Incubate submerged membrane @ room temp 4hr, or 4C ON. Wash 2x 1X PBST. Wash 2x dH2O. 1ary incubation / 2ary incubation / wash / ECL.

Heat dependent antigens

Certain antigens (caveolin-1) have been reported to be heat sensitive in SDS-PAGE analysis. Antigen sensitivity may be improved by adjusting lysis heating parameters from

95C for 3-5 minutes

TO

70C for 10 minutes

Transfer tips for high molecular weight proteins

Standard protein transfer1) Perform electrophoresis on 8-15% acrylamide SDS-PAGE gel, according to standard procedures.

2) Transfer proteins onto a nitrocellulose membrane, overnight (approximately 18 hours) at 0.1 Amp.

3) Running Buffer: 50 mM Tris: 6.06 g Tris Base, 380 mM Glycine: 28.5 g Glycine, 1.6 mM EDTA: 0.67 g EDTA (tetra Na is the most soluble), 0.1% SDS: 1.0 g SDS

4) Transfer buffer: 50 mM Tris, 380 mM glycine, 0.1% (w/v) SDS, 10% (v/v) methanol. Final pH: 8.3.

Adjustments to consider for high molecular weight proteins

6% acrylamide gel is preferable with high molecular weight proteins. Use 15-20% methanol in transfer buffer. Methanol strips the SDS off of the proteins, which

changes the charge of the proteins. Methanol also causes the gel to shrink. Do not let the gel sit in transfer buffer for prolonged periods prior to transfer. Transfer for 16 hours at 30 Volts, then 1 hour at 100 Volts. Do not exceed 220 mAmps. O.45 um PVDF or nitrocellulose may be effective.

Transfer tips for low molecular weight proteins

Tricine Gels are ideal for protein separation from 2.5-50 kDa.

Tricine Gel Gradient 10-20% 0.1 or 0.05um PVDF or 2 pieces of 0.2 micron PVDF lower transfer time relative to standard transfer time

Page 17: Protein Blotting

Tricine GelTricine ions migrate ahead of the smallest proteins. This improves the resolution of low molecular weight proteins. Tricine gels display good transfer efficiency, in a lower pH.

Tricine Gel Recipe

Protein Deglcyosylation

N-link carbohydrate chains (light blue) influence the actual surface contour of membrane bound proteins (green/red) that

harbor them

Glycosylation of proteins and lipids is a post-translational modification important for numerous physiological processes; protein folding, ligand binding, cell surface contour, and influencing cell–cell and cell–matrix interactions.

In order to determine the presence and/or significance of protein glycosylation, the carbohydrate chains may be removed by catalysis.

N-Glycosidase F (PNGase F) is a ~36 kDa amidase that cleaves at the innermost N-acetylglucosamine (GlcNAc) and asparagine residues of high mannose, hybrid, and complex oligosaccharides from N-linked glycoproteins.

PNGase F Endo-O-Glycosidase a-2(3,6,8,9)-Neuraminidase (Sialidase A) ß(1,4)-Galactosidase ß-N-Acetylglycosaminidase

(1) Tarentino, A.L. and Plummer, T.H., Jr. (1994) Methods in Enzymology, 230: 44-47. PMID: 8139511

Deglycosylation kitsEnzymes and reagents combined into convenient to use kits exist in order to remove N-linked and O-linked carbohydrates from glycoproteins, and cleave complex core O-linked carbohydrates (including

Page 18: Protein Blotting

polylactosamine). Deglycosylation can simplify SDS-PAGE by decreasing the potential broad size variation associated with glycoproteins.

New England Biolabs offers a convenient kit Cat#P0704S.

Sigma Aldrich offers the PNGase F enzyme Cat#P7367.

Troubleshooting

Diluting the primary antibody further out in 5% milk TTBS would reduce the overall signal and improve the clarity of the

major bands. Adjusting the exposure time & additional shake rinse after 2ary incubation will help

Western Blot Optimization

Blocking: To block nonspecific sites on the membrane, incubate the membrane with blocking buffer (5% milk TTBS) prior to primary antibody incubation. 2 hour room temperature blocking is sufficient for the membrane to absorb proteins and minimize noise.

Primary antibody: Use a diluent containing a blocking protein (5% milk TTBS or 1%milk/1%BSA TTBS) to dilute the primary antibody.

Secondary antibody: Use a diluent containing a blocking protein (5% milk TTBS or 1%milk/1%BSA TTBS) to dilute the primary antibody. Performing a "Secondary control" control blot where the primary antibody is purposely omitted is useful for identifying whether nonspecific signals are due to the secondary antibody conjugate or the primary antibody.

Washes: All wash steps are critical for reducing general background signal and nonspecific binding to discrete bands. If high background is a problem, the number, length and composition of the washes can all be increased.

General: Handle membranes carefully with forceps to minimize problems with nonspecific signals. Wear gloves for all steps to prevent hand contact with film, membranes or detection reagents.

PVDF versus Nitrocellulose

The western blotting procedures are the same for PVDF or nitrocellulose, however the handling of these membranes are different prior to- and during- transfer of proteins from the SDS-PAGE gel to the membrane.

NitrocelluloseNitrocellulose exhibits the highest sensitivity with very low backgrounds in all transfers, especially in protein blotting. Unlike PVDF, nitrocellulose wets out naturally, does not require methanol, and will not turn hydrophobic during the transfer process. Nitrocellulose is very easily blocked and does not need

Page 19: Protein Blotting

the many blocking steps required with PVDF. Protocols for Western Blotting with PVDF and Nitrocellulose are the same with a few exceptions.

PVDFPolyvinylidene Fluoride is hydrophobic and should be prewet in methanol before it is used.

Wet the membrane in 100% methanol for 15 seconds. The membrane should uniformly change from opaque to semi-transparent.

Place the semi-transparent membrane into ultrapure water and soak for 5 minutes. Place the membrane into transfer buffer and soak for at least 5 minutes. Transfer proteins to the smooth/reflective side that has more shine

Another change to note is that the SDS tolerances are not equivalent for PVDF and Nitrocellulose. The binding of protein to PVDF is much more sensitive to SDS levels. Too much SDS can inhibit the protein's ability to bind to the PVDF and can, in fact, help proteins already bound to the membrane to slip off. SDS levels should never exceed 0.05% for PVDF.

Membrane Stripping

Griffin:Membrane_Stripping_and_Reprobing

Stability of reagents

Ammonium persulphate/persulfate (APS) is a very reactive compound (used for radical generation in the polymerisation process). It is prone to degradation and loss of activity which will increase your polymerisation times. Make small aliquots, keep in the cold. [1]

TEMED, the catalyst of polymerisation, is also prone to oxidation especially if mixed with water which it attracts if the container is left open. Seal quickly, keep in cold place. [2] [3]

Toxicity of reagents

acrylamide/bis-acrylmide is a neurotoxin. Handle with precaution. [4] TEMED is very corrosive. Most likely exposure in the lab due to inhalation. Reseal the container

as quickly as possible. [5]

Introduction to protein blotting :

Protein blotting is a powerful and important procedure for the immunodetection of proteins following electrophoresis, particularly proteins that are of low abundance. Since the inception of the protocol for protein transfer from an electrophoresed gel to a membrane in 1979, protein blotting has evolved greatly. The scientific community is now confronted with a variety of ways and means to carry out this transfer.

Western Blot: Technique, Theory, and Trouble ShootingTahrin Mahmood and Ping-Chang Yang

Page 20: Protein Blotting

Author information   ►  Copyright and License information   ►

This article has been cited by other articles in PMC.

AbstractGo to:

Introduction

Western blot is often used in research to separate and identify proteins. In this technique a mixture of proteins is separated based on molecular weight, and thus by type, through gel electrophoresis. These results are then transferred to a membrane producing a band for each protein. The membrane is then incubated with labels antibodies specific to the protein of interest.

The unbound antibody is washed off leaving only the bound antibody to the protein of interest. The bound antibodies are then detected by developing the film. As the antibodies only bind to the protein of interest, only one band should be visible. The thickness of the band corresponds to the amount of protein present; thus doing a standard can indicate the amount of protein present. The paper will first describe the protocol for western blot, accompanied by pictures to help the reader and theory to rationalize the protocol. This will be followed by the theoretical explanation of the procedure, and in the later section, troubleshooting tips for common problems.

Technique

Cell lysis to extract protein

Protein can be extracted from different kind of samples, such as tissue or cells. Below is the protocol to extract proteins from adherent cells.

Adherent cells:

1. Wash cells in the tissue culture flask or dish by adding cold phosphate buffered saline (PBS) and rocking gently. Discard PBS. (Tip: Keep tissue culture dish on ice throughout).

2. Add PBS and use a cell scraper to dislodge the cells. Pipette the mixture into microcentrifuge tubes.

3. Centrifuge at 1500 RPM for 5 minutes and discard the supernatant.4. Add 180 μL of ice cold cell lysis buffer with 20 μL fresh protease inhibitor cocktail. (Tip: If

protein concentration is not high enough at the end, it is advised to repeat the procedure with a higher proportion of protease inhibitor cocktail).

5. Incubate for 30 minutes on ice, and then clarify the lysate by spinning for 10 minutes at 12,000 RPM, at 4°C.

6. Transfer supernatant (or protein mix) to a fresh tube and store on ice or frozen at -20°C or -80°C.

7. Measure the concentration of protein using a spectrophotometer.

Page 21: Protein Blotting

Sample preparation

1.

determine the volume of protein extract to ensure 50 μg in each well.

2. Add 5 μL sample buffer to the sample, and make the volume in each lane equalized using double distilled H2O (dd H2O). Mix well. (Tip: Total volume of 15 μL per lane is suggested).

3. Heat the samples with dry plate for 5 minutes at 100°C.

Gel preparation

1. After preparing the 10% stacking gel solution, assemble the rack for gel solidification [Figure 1]. (Tip: 10% AP and TEMED solidify the solution; therefore, both gels can be prepared at the same time, if the abovementioned reagents are not added until the end).

Figure 1

Assembled rack for gel solidification

2. Add stacking gel solution carefully until the level is equal to the green bar holding the glass plates [Figure 2]. Add H2O to the top. Wait for 15–30 minutes until the gel turning solidified. (Tip: Using a suction pipette can make the process of adding the gel to the glass plate easier).

Figure 2

Add gel solution using a transfer pipette

Page 22: Protein Blotting

3. Overlay the stacking gel with the separating gel, after removing the water. (Tip: It is better to tilt the apparatus and use a paper towel to remove the water).

4. Insert the comb, ensuring that there are no air bubbles.5. Wait until the gel is solidified. (Tip: Solidification can be easily checked by leaving some gel

solution in a tube).

Electrophoresis

1. Pour the running buffer into the electrophorator [Figure 3].

Figure 3

Add running buffer to the electrophorator

2. Place gel inside the electrophorator and connect to a power supply. (Tip: When connecting to the power source always connect red to red, and black to black).

3. Make sure buffer covers the gel completely, and remove the comb carefully.4. Load marker (6 μL) followed by samples (15 μL) in to each well [Figure 4].

Figure 4

Add samples and molecular marker to the gel, after removing the combs

5. Run the gel with low voltage (60 V) for separating gel; use higher voltage (140 V) for stacking gel [Figure[Figure5a 5a and andb b ].

Figure 5

(a) Samples running through the stacking gel (lower voltage). (b): Samples running through the separating gel (higher voltage)

Page 23: Protein Blotting

6. Run the gel for approximately an hour, or until the dye front runs off the bottom of the gel [Figure 6].

Figure 6

Run the gel to the bottom of the electrophorator

Electrotransfer

1. Cut 6 filter sheets to fit the measurement of the gel, and one polyvinylidene fluoride (PDVF) membrane with the same dimensions.

2. Wet the sponge and filter paper in transfer buffer, and wet the PDVF membrane in methanol.

3. Separate glass plates and retrieve the gel.4. Create a transfer sandwich as follows:

Sponge

3 Filter Papers

Gel PVDF

3 Filter Papers

(Tip: Ensure there are no air bubbles between the gel and PVDF membrane, and squeeze out extra liquid).

5. Relocate the sandwich to the transfer apparatus, which should be placed on ice to maintain 4°C. Add transfer buffer to the apparatus, and ensure that the sandwich is covered with the buffer. Place electrodes on top of the sandwich, ensuring that the PVDF membrane is between the gel and a positive electrode [Figure 7].

Figure 7

Transfer should be done on ice

Page 24: Protein Blotting

6. Transfer for 90 minutes [Figure 8]. (Tip: The running time should be proportional to the thickness of the gel, so this may be reduced to 45 minutes for 0.75 mm gels).

Figure 8

Membrane after transfer

Blocking and antibody incubation

1. Block the membrane with 5% skim milk in TBST* for 1 hour.2. Add primary antibody in 5% bovine serum albumin ( BSA) and incubate overnight in 4°C on a

shaker [Figure 9].

Figure 9

Use a shaker to incubate the membrane with antibody

3. Wash the membrane with TBST for 5 minutes. Do this 3 times. (Tip: All washing and antibody incubation steps should be done on a shaker at room temperature to ensure even agitation).

4. Add secondary antibody in 5% skim milk in TBST, and incubate for 1 hour.5. Wash the membrane with TBST for 5 minutes. Do this 3 times6. Prepare ECL mix (following the proportion of solution A and B provided by the

manufacturer). Incubate the membrane for 1–2 minutes [Figure 10]. (Tip: Use a 1000 μL pipette to ensure that ECL covers the top and bottom of the membrane).

Figure 10

Incubate the membrane with ECL mix using a 1000 μL pipette to help the process

Page 25: Protein Blotting

7. Visualize the result in the dark room [Figure 11]. (Tip: If the background is too strong, reduce exposure time).

Figure 11

Use the cassette to expose the membrane in the dark room

Recipe

1. Dissolve the following in 800 ml of distilled H2O 8.8 g of NaCl 0.2g of KCl 3g of Tris base

2. Add 500ul of Tween-203. Adjust the pH to 7.44. Add distilled H2O to 1L5. Sterilize by filtration or autoclaving

Theory

Sample preparation

Cell lysates are the most common form of sample used for western blot. Protein extraction attempts to collect all the proteins in the cell cytosol. This should be done in a cold temperature with protease inhibitors to prevent denaturing of the proteins. Since tissue sample display a higher degree of structure, mechanical invention, such as homogenization, or sonication is needed to extract the proteins.

After extracting the protein, it is very important to have a good idea of the extract's concentration. This eventually allows the researcher to ensure that the samples are being compared on an equivalent basis. Protein concentration is often measured using a spectrophotometer. Using this concentration allows to measure the mass of the protein that is being loaded into each well by the relationship between concentration, mass, and volume.

After determining the appropriate volume of the sample, it is diluted into a loading buffer, which contains glycerol so that the samples sink easily into the wells of the gel. A tracking dye (bromophenol blue) is also present in the buffer allowing the researcher to see how far the separation has progressed. The sample is heated after being diluted into a loading buffer, in order to denature the higher order structure, while retaining sulfide bridges. Denaturing the high structure ensures that the negative charge of amino acids is not neutralized, enabling the protein to move in an electric field (applied during electrotransfer).

Page 26: Protein Blotting

It is also very important to have positive and negative controls for the sample. For a positive control a known source of target protein, such as purified protein or a control lysate is used. This helps to confirm the identity of the protein, and the activity of the antibody. A negative control is a null cell line, such as β-actin, is used as well to confirm that the staining is not nonspecific.

Gel electrophoresis

Western blot uses two different types of agarose gel: stacking and separating gel. The higher, stacking gel is slightly acidic (pH 6.8) and has a lower acrylamide concentration making a porous gel, which separates protein poorly but allows them to form thin, sharply defined bands. The lower gel, called the separating, or resolving gel, is basic (pH 8.8), and has a higher polyacrylamide content, making the gel's pores narrower. Protein is thus separated by their size more so in this gel, as the smaller proteins to travel more easily, and hence rapidly, than larger proteins.

The proteins when loaded on the gel have a negative charge, as they have been denatured by heating, and will travel toward the positive electrode when a voltage is applied. Gels are usually made by pouring them between two glass or plastic plates, using the solution described in the protocol section. The samples and a marker are loaded into the wells, and the empty wells are loaded with sample buffer. The gel is then connected to the power supply and allowed to run. The voltage is very important, as a high voltage can overheat and distort the bands.

Blotting

After separating the protein mixture, it is transferred to a membrane. The transfer is done using an electric field oriented perpendicular to the surface of the gel, causing proteins to move out of the gel and onto the membrane. The membrane is placed between the gel surface and the positive electrode in a sandwich. The sandwich includes a fiber pad (sponge) at each end, and filter papers to protect the gel and blotting membrane [Figure 12]. Here two things are very important: (1) the close contact of gel and membrane to ensure a clear image and (2) the placement of the membrane between the gel and the positive electrode. The membrane must be placed as such, so that the negatively charged proteins can migrate from the gel to the membrane. This type of transfer is called electrophoretic transfer, and can be done in semi-dry or wet conditions. Wet conditions are usually more reliable as it is less likely to dry out the gel, and is preferred for larger proteins.

Figure 12

Assembly of a sandwich in western Blot

Page 27: Protein Blotting

The membrane, the solid support, is an essential part of this process. There are two types of membrane: nitrocellulose and PVDF. Nitrocellulose is used for its high affinity for protein and its retention abilities. However, it is brittle, and does not allow the membrane to be used for reprobing. In this regard, PVDF membranes provide better mechanical support and allow the blot to be reprobed and stored. However, the background is higher in the PVDF membranes and therefore, washing carefully is very important.

Washing, blocking and antibody incubation

Blocking is a very important step of western blotting, as it prevents antibodies from binding to the membrane nonspecifically. Blocking is often made with 5% BSA or nonfat dried milk diluted in TBST to reduce the background.

Nonfat dried milk is often preferred as it is inexpensive and widely available. However, milk proteins are not compatible with all detection labels, so care must be taken to choose the appropriate blocking solution. For example, BSA blocking solutions are preferred with biotin and AP antibody labels, and antiphosphoprotein antibodies, since milk contains casein, which is itself a phosphoprotein and biotin, thus interfering with the assay results. It is often a good strategy to incubate the primary antibody with BSA since it is usually needed in higher amounts than the secondary antibody. Putting it in BSA solution allows the antibody to be reused, if the blot does not give good result.

The concentration of the antibody depends on the instruction by the manufacturer. The antibody can be diluted in a wash buffer, such as PBS or TBST. Washing is very important as it minimized background and removes unbound antibody. However, the membrane should not be left to wash for a really long time, as it can also reduce the signal.

The membrane is then detected using the label antibody, usually with an enzyme such as horseradish peroxidase (HRP), which is detected by the signal it produces corresponding to the position of the target protein. This signal is captured on a film which is usually developed in a dark room.

Quantification

It is very important to be aware that the data produced with a western blot is typically considered to be semi-quantitative. This is because it provides a relative comparison of protein levels, but not an absolute measure of quantity. There are two reasons for this; first, there are variations in loading and transfer rates between the samples in separate lanes which are different on separate blots. These differences will need to be standardized before a more precise comparison can be made. Second, the signal generated by detection is not linear across the concentration range of samples. Thus, since the signal produced is not linear, it should not be used to model the concentration.

Troubleshooting

Even though the procedure for western blot is simple, many problems can arise, leading to unexpected results. The problem can be grouped into five categories: (1) unusual or

Page 28: Protein Blotting

unexpected bands, (2) no bands, (3) faint bands or weak signal, (4) high background on the blot, and (5) patchy or uneven spots on the blot.

Unusual or unexpected bands can be due to protease degradation, which produces bands at unexpected positions. In this case it is advisable to use a fresh sample which had been kept on ice or alter the antibody. If the protein seems to be in too high of a position, then reheating the sample can help to break the quaternary protein structure. Similarly, blurry bands are often caused by high voltage or air bubbles present during transfer. In this case, it should be ensured that the gel is run at a lower voltage, and that the transfer sandwich is prepared properly. In addition, changing the running buffer can also help the problem. Nonflat bands can be the result of too fast of a travel through the gel, due to low resistance. To fix this the gel should be optimized to fit the sample. Finally, white (negative) bands on the film are due to too much protein or antibody.

Another problem: no bands can also arise due to many reasons related to antibody, antigen, or buffer used. If an improper antibody is used, either primary or secondary, the band will not show. In addition, the concentration of the antibody should be appropriate as well; if the concentration is too low, the signal may not be visible. It is important to remember that some antibodies are not to be used for western blot. Another reason for no visible bands is the lowest concentration or absence of the antigen. In this case, antigen from another source can be used to confirm whether the problem lies with the sample or with other elements, such as the antibody. Moreover, prolonged washing can also decrease the signal. Buffers can also contribute to the problem. It should be ensured that buffers like the transfer buffer, TBST, running buffer and ECL are all new and noncontaminated. If the buffers are contaminated with sodium azide, it can inactivate HRP.

Similarly, weak signals can be caused by low concentration of antibody or antigen. Increasing exposure time can also help to make the band clearer. Another reason could be nonfat dry milk masking the antigen. In this case use BSA or decrease the amount of milk used.

High background is often caused by too high concentration of the antibody, which can bind to PVDF membranes. Another problem could be the buffers, which may be too old. Increasing the washing time can also help to decrease the background. Additionally, too high of an exposure can also lead to this problem. Therefore, it is advisable to check different exposure times to achieve an optimum time.

Patchy and uneven spots on the blot are usually caused by improper transfer. If there are air bubbles trapped between the gel and the membrane, it will appear darker on the film. It is also important to use a shaker for all incubation, so that there is no uneven agitation during the incubation. Once again, washing is of utmost importance as well to wash the background. This problem can also be caused by antibodies binding to the blocking agents; in this case another blocking agent should be tried. Filtering the blocking agent can also help to remove some contaminants. Finally, this problem can also be caused by aggregation of the secondary antibody; in this case, the secondary antibody should be centrifuged and filtered to remove the aggregated.

Page 29: Protein Blotting

Go to:

Conclusion

Western blot is a technique that is very useful for protein detection as it allows the user to quantify the protein expression as well. This paper covered the protocol, the theory behind that protocol, and some troubleshooting techniques. Western blot can be seen as an intricate balance, as the researcher attempts to get a nonspecific, yet strong signal.

Western Blot Principle / Western Blotting Principle

Western blot is an important technique used in cell and molecular biology. By using a western blot, researchers are able to identify specific proteins from a complex mixture of proteins extracted from cells. The basic principle of western blot is to use three elements to accomplish this task: (1) separation by size, (2) transfer to a solid support, and (3) marking target protein using a proper primary and secondary antibody to visualize.In this technique a mixture of proteins is separated based on molecular weight, and thus by type, through gel electrophoresis. These results are then transferred to a membrane producing a band for each protein by western blot transfer. The membrane is then incubated with labels antibodies specific to the protein of interest.The unbound antibody is washed off leaving only the bound antibody to the protein of interest. The bound antibodies are then detected by developing the film in the procedure of western blot detection. As the antibodies only bind to the protein of interest, only one band should be visible. The thickness of the band corresponds to the amount of protein present; thus doing a standard can indicate the amount of protein present.Content

Sample Preparation Principle

SDS-PAGE Principle

Western Blot Transfer Principle

Blocking and Antibody Incubation Principle

Western Blot Quantification Principle

Western Blot Principle: Sample Preparation PrincipleCell lysates are the most common form of sample used for western blot. Protein extraction attempts to collect all the proteins in the cell cytosol. This should be done in a cold temperature with protease inhibitors to prevent denaturing of the proteins. Since tissue sample display a higher

Page 30: Protein Blotting

degree of structure, mechanical invention, such as homogenization, or sonication is needed to extract the proteins.After extracting the protein, it is very important to have a good idea of the extract's concentration. This eventually allows the researcher to ensure that the samples are being compared on an equivalent basis. Protein concentration is often measured using a spectrophotometer. Using this concentration allows to measure the mass of the protein that is being loaded into each well by the relationship between concentration, mass, and volume.After determining the appropriate volume of the sample, it is diluted into a loading buffer, which contains glycerol so that the samples sink easily into the wells of the gel. A tracking dye (bromophenol blue) is also present in the buffer allowing the researcher to see how far the separation has progressed. The sample is heated after being diluted into a loading buffer, in order to denature the higher order structure, while retaining sulfide bridges. Denaturing the high structure ensures that the negative charge of amino acids is not neutralized, enabling the protein to move in an electric field (applied during electrotransfer).It is also very important to have positive and negative controls for the sample. For a positive control a known source of target protein, such as purified protein or a cell lysate is used. This helps to confirm the identity of the protein, and the activity of the antibody. A negative control is a null cell line, such as β-actin, is used as well to confirm that the staining is not nonspecific.

Western Blot Principle: SDS-PAGE PrincipleWestern blot uses two different types of agarose gel: stacking and separating gel. The higher, stacking gel is slightly acidic (pH 6.8) and has a lower acrylamide concentration making a porous gel, which separates protein poorly but allows them to form thin, sharply defined bands. The lower gel, called the separating, or resolving gel, is basic (pH 8.8), and has a higher polyacrylamide content, making the gel's pores narrower. Protein is thus separated by their size more so in this gel, as the smaller proteins to travel more easily, and hence rapidly, than larger proteins.The proteins when loaded on the gel have a negative charge, as they have been denatured by heating, and will travel toward the positive electrode when a voltage is applied. Gels are usually made by pouring them between two glass or plastic plates, using the solution described in the protocol section. The samples and a marker are loaded into the wells, and the empty wells are loaded with sample buffer. The gel is then connected to the power supply and allowed to run. The voltage is very important, as a high voltage can overheat and distort the bands.

Western Blot Principle: Western Blot Transfer PrincipleAfter separating the protein mixture, it is transferred to a membrane. The transfer is done using an electric field oriented perpendicular to the surface of the gel, causing proteins to move out of the gel and onto the membrane. The membrane is placed between the gel surface and the positive electrode in a sandwich. The sandwich includes a fiber pad (sponge) at each end, and filter papers to protect the gel and blotting membrane. Here two things are very important: (1) the close contact of gel and membrane to ensure a clear image and (2) the placement of the membrane between the gel and the positive electrode. The membrane must be placed as such, so that the negatively charged proteins can migrate from the gel to the membrane. This type of

Page 31: Protein Blotting

transfer is called electrophoretic transfer, and can be done in semi-dry or wet conditions. Wet conditions are usually more reliable as it is less likely to dry out the gel, and is preferred for larger proteins.The membrane, the solid support, is an essential part of this process. There are two types of membrane: nitrocellulose and PVDF. Nitrocellulose is used for its high affinity for protein and its retention abilities. However, it is brittle, and does not allow the membrane to be used for reprobing. In this regard, PVDF membranes provide better mechanical support and allow the blot to be reprobed and stored. However, the background is higher in the PVDF membranes and therefore, washing carefully is very important.

Western Blot Principle: Blocking   and Antibody Incubation PrincipleBlocking is a very important step of western blotting, as it prevents antibodies from binding to the membrane nonspecifically. Blocking is often made with 5% BSA or nonfat dried milk diluted in TBST to reduce the background.Nonfat dried milk is often preferred as it is inexpensive and widely available. However, milk proteins are not compatible with all detection labels, so care must be taken to choose the appropriate blocking solution. For example, BSA blocking solutions are preferred with biotin and AP antibody labels, and antiphosphoprotein antibodies, since milk contains casein, which is itself a phosphoprotein and biotin, thus interfering with the assay results. It is often a good strategy to incubate the primary antibody with BSA since it is usually needed in higher amounts than the secondary antibody. Putting it in BSA solution allows the antibody to be reused, if the blot does not give good result.The concentration of the antibody depends on the instruction by the manufacturer. The antibody can be diluted in a wash buffer, such as PBS or TBST. Washing is very important as it minimized background and removes unbound antibody. However, the membrane should not be left to wash for a really long time, as it can also reduce the signal.The membrane is then detected using the label antibody, usually with an enzyme such as horseradish peroxidase (HRP), which is detected by the signal it produces corresponding to the position of the target protein. This signal is captured on a film which is usually developed in a dark room.

Western Blot Principle: Western Blot Quantification PrincipleIt is very important to be aware that the data produced with a western blot is typically considered to be semi-quantitative. This is because it provides a relative comparison of protein levels, but not an absolute measure of quantity. There are two reasons for this; first, there are variations in loading and transfer rates between the samples in separate lanes which are different on separate blots. These differences will need to be standardized before a more precise comparison can be made. Second, the signal generated by detection is not linear across the concentration range of samples. Thus, since the signal produced is not linear, it should not be used to model the concentration

Page 32: Protein Blotting

Western blotting(also called protein immunoblotting because an antibody is used to specifically detect its

antigen) is a widely accepted analytical technique used to detect specific proteins in the given sample. It

uses SDS-polyacrylamide gel electrophoresis(SDS-PAGE) to separate various proteins contained in the

given sample(e.g. to separate native proteins by 3-D structure or denatured proteins by the length of the

polypeptide). The separated proteins are then transferred or blotted onto a matrix( generally nitrocellulose

or PVDF membrane), where they are stained with antibodies(used as a probe) specific to the target protein.

By analyzing location and intensity of the specific reaction, expression details of the target proteins in the

given cells or tissue homogenate could be obtained. Western blotting could detect target protein which is

as low as 1ng due to high resolution of the gel electrophoresis and strong specificity and high sensitivity of

the immunoassay. This method is used in the fields of molecular biology, biochemistry, immunogenetics

and other molecular biology disciplines.

1. PrincipleWestern blotting usually involves two major processes, namely, SDS-polyacrylamide gel electrophoresis

and protein blotting and testing.SDS-polyacrylamide gel electrophoresis(SDS-PAGE)Electrophoresis separation describes a phenomenon that charged particles move towards opposite electrode

under the influence of electric field. It is used to separate proteins according to their electrophoretic

mobility which depends on charge, molecule size and structure of the proteins. Polyacrylamide gel(PAG)

is a three-dimensional mesh networks polymer composed of acrylamide and a cross-linker(methylene

bisacrylamide) under the catalyzation of ammonium persulfate. PAG is a versatile supporting matrix due to

its stable hydrophily and little adsorption and electroosmosis effect provided by its neutrally charged

nature.(It possesses several electrophoretically desirable features that make it a versatile medium. It is a

synthetic, thermo-stable, transparent, strong, chemically relatively inert gel, and can be prepared with a

wide range of average pore sizes.) In the presence of SDS, electrophoretic mobility is mainly based on

molecule weight instead of on charge and size of the proteins. SDS is an anionic detergent which could

break hydrogen bond within and between molecules to unfold proteins and break up secondary and tertiary

structures as denaturing agent and hydrotropy agent. Strong reducing agents such as mercaptoethanol and

Dithiothreitol(DTT) could disrupt disulfide linkages between cysteine residues. SDS and reducing agents

are applied to protein sample to linearize proteins and to impart a negative charge to linearized proteins. In

most proteins, the binding of SDS to the polypeptide chain imparts an even distribution of charge per unit

mass, thereby the intrinsic charges of polypeptides becomes negligible when compared to the negative

charges contributed by SDS. This new negative charge is significantly greater than the original charge of

that protein. The electrostatic repulsion that is created by binding of SDS causes proteins to unfold into a

rod-like shape thereby eliminating differences in shape as a factor for separation in the gel. Minor axis of

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all rods, the SDS-protein subunit compound are nearly the same, about 1.8nm. And the length of major

axis is in proportion to molecular weight of the protein subunit. Thus electrophoretic mobility of the SDS-

protein subunit compound is based on molecular weight, eliminating the influence imposed by size and

charge. The sample to be analyzed is mixed with SDS. And the mixed samples are subsquently treated by

related solution. Heating the samples to at least 60°C further promotes protein denaturation and

depolymerization, helping SDS to bind and enabling the rod-shape formation and negative charge

adherence. A bromophenol blue dye may be added to the protein solution to allow the experimenter to

track the progress of the protein solution through the gel during the electrophoretic run. An appropriate

amount of glycerol is added to increase density and accelerate the migration of sample solution.

A buffer system with different pH values is applied in gel electrophoresis process. A very widespread

discontinuous buffer system is the tris-glycine or "Laemmli" system that stacks at a pH of 6.8 and resolves

at a pH of ~8.3-9.0. A drawback of this system is that these pH values may promote disulfide bond

formation between cysteine residues in the proteins because the pKa of cysteine ranges from 8-9 and

because reducing agent present in the loading buffer doesn't co-migrate with the proteins. Recent advances

in buffering technology alleviate this problem by resolving the proteins at a pH well below the pKa of

cysteine (e.g., bis-tris, pH 6.5) and include reducing agents (e.g. sodium bisulfite) that move into the gel

ahead of the proteins to maintain a reducing environment. An additional benefit of using buffers with

lower pH values is that the acrylamide gel is more stable at lower pH values, so the gels can be stored for

long periods of time before use.

As voltage is applied, the anions (and negatively charged sample molecules) migrate toward the positive

electrode (anode) in the lower chamber, the leading ion is Cl¯ ( high mobility and high concentration);

glycinate is the trailing ion (low mobility and low concentration). SDS-protein particles do not migrate

freely at the border between the Cl¯ of the gel buffer and the Gly¯ of the cathode buffer. Because of the

voltage drop between the Cl- and Glycine-buffers, proteins are compressed (stacked) into micrometer thin

layer-stacking gel layer. In resolving gel layer, proteins with more negative charges per unit migrate faster

than those with less negative charges per unit. That is, proteins with small molecular weight migrate faster

than proteins with large molecular weight. The boundary moves through a pore gradient and the protein

stack gradually disperses due to a frictional resistance increase of the gel matrix. Stacking and unstacking

occurs continuously in the gradient gel, for every protein at a different position.

Polyacrylamide gel electrophoresis (PAGE) is used for separating proteins ranging in size from 5 to 2,000

kDa due to the uniform pore size provided by the polyacrylamide gel. Pore size is controlled by controlling

the concentrations of acrylamide and bis-acrylamide powder used in creating a gel. Typically resolving

gels are made in 5%, 8%, 10%, 12% or 15%. Stacking gel (5%) is poured on top of the resolving gel and a

gel comb (which forms the wells and defines the lanes where proteins, sample buffer and ladders will be

placed) is inserted. The percentage chosen depends on the size of the protein that one wishes to identify or

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probe in the sample. The smaller the known weight, the higher the percentage that should be used.

Changes on the buffer system of the gel can help to further resolve proteins of very small sizes.Range of molecular weight(KD) Concentration of gel(%)

<10 15

10-30 12

30-100 10

100-500 8

>500 5

Western blotting and detectionFive steps are involved in western blotting and detection assay, namely, transfer, blocking, primary

antibody incubation, secondary antibody incubation and protein detection and analysis.

Transfer

Proteins are moved from within the gel onto a membrane made of nitrocellulose(NC) or polyvinylidene

difluoride(PVDF). Without pre-activation, proteins combine with nitrocellulose membrane based on

hydrophobic interaction, thereby having slight effect on protein activities. Besides, nitrocellulose

membrane produces little non-specific staining. It is cheap and ease to use. However, it is easy to erase

small molecular proteins while washing. It is fragile and has poor toughness. With high affinity, the PVDF

membrane needs to be sunk in methanol before use to activate positive charge groups on the membrane,

promoting combination with negative charged proteins. Specific NC membrane with different pores should

be applied according to the molecular weight of transferred proteins due to the smaller the pore of

membrane the tighter the combination between membrane and small molecular weight proteins. NC

membranes of 0.45 µm and of 0.2 µm are used most. The size of 0.45 µm should be applied for proteins

with molecular weight over 20KD while the size of 0.2 µm will be chosen for those below 20KD. PVDF

membrane is best for the detection of small molecular weight proteins due to its higher sensitivity,

resolution as well as affinity than normal membrane. 

Transfer methods that are used most for proteins are semi-dry transfer and wet transfer. Semi-dry transfer

describes the method that Gel-Membrane-Filter sandwich is placed between filters loaded with transfer

buffer. The transfer process is based on current conduction produced by the transfer buffer. Semi-dry

transfer takes little time with high efficiency as electric current works directly on membrane and gel. While

applying wet transfer, the Gel-Membrane-Filter sandwich is placed in the transfer tank, suspending in

transfer buffer vertically. Proteins transfer from the gel to the membrane under the control of high intensity

electric field produced by electrode plate paralleled to the sandwich. While prolonging time to an

appropriate extend, proteins could be transferred more effectively. Proteins within several gels could be

transferred.

Blocking

In a western blot, it is important to block the unreacted sites on the membrane to reduce the amount of

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nonspecific binding of proteins during subsequent steps in the assay using inert protein or nonionic

detergent. Blocking buffers should block all unreacted sites. And Blocking buffers should not replace

target protein on the membrane, not bind epitope on the target protein and not cross react with antibody or

detection reagents. The most typical blockers are BSA, nonfat dry milk, casein, gelatin and Tween-20.

TBS and/or PBS are the most commonly used buffers. 

Inertia protein BSA, nonfat dry milk, casein, gelatin or nonionic detergent Tween-20 reduce nonspecific

binding by blocking unreacted sites. Retaining protein structure, Tween-20 can reduce breakup to original

interaction among proteins while is used for protein emulsification.

Nonfat dry milk is the most economic choice

Avoid using nonfat dry milk as a blocking reagent for blots with biotin conjugated antibody because milk

contains variable amounts of glycoprotein and biotin.

BSA is appropriate for blots with phosphorylated protein as target. Phosphatase contained in nonfat dry

milk leads to dephosphorylation of phosphorylated protein on the membrane while phosphoryltion specific

antibody is used to identify phosphorylated protein. And nonfat dry milk is improper for blots which rely

on alkaline phosphatase system.

Avoid adding NaN3 into blocking reagent for blots that base on HRP system because NaN3 is enabled to

inactivate HRP.

Casein is recommended for blots with alkaline phosphatase conjugated secondary antibody. TBS buffer

instead of PBS buffer should be chosen because PBS interferes alkaline phosphatase.

Primary antibody incubation

After blocking, primary antibody specific to target protein is incubated with the membrane. And the

primary antibody binds to target protein on the membrane.

In western blot, primary antibody should be validated before use. The choice of a primary antibody

depends on the antigen to be detected. Both polyclonal and monoclonal antibodies work well for western

blot. Monoclonal antibodies recognize single specific antigenic epitope. Thus, they have higher specificity

resulting in lower background. Blot results will be influenced if the target epitope is destroyed. Polyclonal

antibodies recognize more epitopes and they often have higher affinity. Blot results will be stable even

though a few epitopes are destroyed.

Secondary antibody incubation

After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to specific

enzyme conjugated secondary antibody. And the secondary antibody binds to the primary antibody which

has reacted with target protein

The most popular secondary antibodies are anti-mouse and anti-rabbit immune globulin since the host

species for primary antibodies are mainly mouse and rabbit. Goat is used widely to raise anti-mouse and

anti-rabbit polyclonal antibodies. Thus, goat anti-mouse and goat anti-rabbit immune globulin are the most

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commonly used secondary antibodies.The choice of secondary antibody depends upon the species of

animal in which the primary antibody was raised. For example, if the primary antibody is a mouse

monoclonal antibody, the secondary antibody must be an anti-mouse antibody. If the primary antibody is a

rabbit polyclonal antibody, the secondary antibody must be an anti-rabbit antibody.

Protein detection(color development) and analysis

1. Protein detection(color development)

A substrate reacts with the enzyme that is bound to the secondary antibody to generate

colored substance, namely, visible protein bands. The target protein levels in cells or

tissues are evaluated though densitometry and location of the visible protein bands.

Alkaline phosphatase (AP) and horseradish peroxidase (HRP) are the two enzymes that

are used extensively. Functioned by Alkaline phosphatase (AP) catalyzation, a colorless

substrate BCIP will be converted to a blue product. In the presence of H2O2, 3-amino-9-

ethyl carbazole and 4-chlorine naphthol will be oxidized into brown substance and blue

product respectively under the catalyzation of HRP. Enhanced chemiluminescence is

another method that employs HPR detection. Using HRP as the enzyme label,

luminescent substance luminol will be oxidized by H2O2 and will luminesce. Moreover,

enhancers in this substrate will enable a 1000-fold increase in light intensity. HRP will be

detected when the blot is sensitized on photographic film.

2. Analysis 

a. Control design

Proper control design is essential to western blot. It will guarantee accurate and specific

test result by identifying various problems quickly and precisely.

Control types:

Positive control: A lysate from a cell line or tissue sample known to express the protein

you are detecting. Positive control is designed to verify working efficiency of the

antibodies. 

Negative control: A lysate from a cell line or tissue sample known not to express the

protein you are detecting. Negative control is to check antibody specificity. Nonspecific

binding and false positive result will be identified. 

Secondary antibody control(No primary antibody control): The primary antibody is

not added to the membrane. Only secondary antibody is added. This is to check secondary

antibody specificity. Nonspecific binding and false positive result caused by secondary

antibody will be indicated. 

Blank control: Both primary and secondary antibody are not added to membrane. This is

to check membrane nature and blocking effect. 

Loading control: Loading control is used to check sample quality and the performance

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of secondary antibody system.

Loading controls are antibodies to "house-keeping proteins", or proteins that are expressed at equivalent

levels in almost all tissues and cells. Loading controls are required to check that the lanes in your gel have

been evenly loaded with sample, especially when a comparison must be made between the expression

levels of a protein in different samples. They are also useful to check for even transfer from the gel to the

membrane across the whole gel. Where even loading or transfer have not occurred, the loading control

bands can be used to quantify the protein amounts in each lane. For publication-quality work, use of a

loading control is absolutely essential.Loading control Molecular weight(KD) Sample type

Beta-Actin 43KD Whole Cell/cytoplasmic

GAPDH 30-40KD Whole Cell/cytoplasmic

Tubulin 55KD Whole Cell/cytoplasmic

VCDA1/Porin 31KD Mitochondrial

COXIV 16KD Mitochondrial

Lamin B1 16KD Nuclear(Not suitable for samples where the nuclearenvelope is removed.)

TBP 38KD Nuclear(Not suitable for samples where DNA is removed)

Beta-Actin antibodyBoster ECL western blot substrate

GAPDH antibodyBoster ECL western blot substrate

2. Western blotting diagram

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