WESTERN BLOTTING
Good afternoon Dr. Villordon and classmates. I am Chonnie Mae Pis-an and I will be
discussing Western Blotting.
To get us started, I will be defining the terms "blotting" and "SDS-PAGE".
 Blotting refers to the transfer of biological samples from a gel to a membrane and
their subsequent detection on the surface of the membrane.
 Sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) is
commonly used to obtain high resolution separation of complex mixtures of
proteins. The method initially denatures the proteins that will undergo
electrophoresis.
Aside from Western Blotting, other "blot protocols" also exist such as the Northern and
Southern Blot. The table shows their similarities and differences.
[ayaw na basaha ning table, daritso na aning paragraphs]
Southern, northern, and western blot protocols are similar, and begin with electrophoretic
separation of protein and nucleic acid fragments on a gel, which are then transferred to a
membrane (nitrocellulose membrane, polyvinylidene difluoride (PVDF) membrane, etc.) where
they are immobilized. This enables radiolabeled or enzymatically labeled antibody or DNA probes
to bind the immobilized target, and the molecules of interest may then be visualized with various
methods. Blotting techniques are selected based on the target molecule: DNA, RNA, or protein.
In summary, the three blot protocols have different target molecules and sample preparations.
The Southern, Northern and Western blot have the same separation technique which is
Electrophoresis. Western blot differs from both the Southern and Northern protocol in terms of
membrane material.
Now, let us redirect our attention to Western Blotting.
DEFINITION
Western blotting
 a well-established analytical technique for detecting, analyzing, and quantifying
a specific protein molecule from among a mixture of proteins associated with a
particular tissue or cell type.
Western blotting typically involves protein separation by gel electrophoresis followed by
transfer to a polyvinylidene difluoride (PVDF) or nitrocellulose membrane. After proteins have
been transferred, they can be stained for visualization and directly identified by N-terminal
sequencing, mass spectrometry or immunodetection. The membrane is exposed to an
antibody specific to the target protein. Binding of the antibody is detected using a radioactive
or chemical tag.
This procedure was named for its similarity to the previously invented method known
as the Southern blot.
PURPOSE
Western blot
General Purposes
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widely used to detect specific protein molecules in complex samples such as tissue
homogenates and cell lysates.
sometimes used to diagnose disease.
can also be used to evaluate the size of a protein of interest, and to measure the
amount of protein expression.
Specific Purposes
The technique enables evaluation of: [only read headings]
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Protein–DNA interactions – Attachment of DNA binding proteins to specific DNA sequences
is vital to the process of transcriptional regulation. Southwestern blotting (similar to western
blotting except the membrane is probed with DNA) can be used to identify transcription
factors in gene regulation studies.
Protein–protein interactions – These are vital for many essential cellular processes.
Detection of protein–protein interactions, using a variation of western blotting known as
far-western blotting, can help to elucidate cellular function and dysfunction.
Post-translational modifications (PTMs) – PTM’s impact protein folding and consequently
function, expanding proteome diversity. However, aberrant PTM’s have been associated
with disease. Consequently, their study is of great interest to researchers and western blot
provides one such tool with which to do this.
Protein isoform detection – Proteins may be expressed in differing isoforms in different
cellular states with varying activities or targets. Alternatively, they may require cleavage to
become activated.
Antibody characterization – When antibodies are produced as tools or therapeutics, their
validation and characterization is vital to ensure correct performance and safety. As an
antibody-based method, western blotting is a useful technique in this process.
Epitope mapping – Understanding how and where antibodies bind their target protein is
valuable for research, diagnostic and therapeutic purposes. There are many tools used
towards this goal and thanks to its specificity, western blotting is one.
Subcellular protein localization – Performing western blot analysis of different cellular
fractions allows the location of target proteins in the cell to be determined. Single-cell
western blotting has offered great insights in this field and overcomes some of the antibody
cross-reactivity issues experienced by other single-cell assays.
BACKGROUND / HISTORY
Western blot experiment, or western blotting was developed in 1979 by Harry Towbin and
his colleagues and they named it “western blot” due to the technique’s similarity to
Southern blotting. It is now a routine technique for protein analysis. It is also called
immunoblotting because an antibody is used to specifically detect its antigen. This
antibody-based technique is used to detect the presence, size and abundance of
specific proteins within a sample. Western blotting can produce qualitative and semiquantitative data about the protein of interest.
WESTERN BLOT PROTOCOL
Overview
For a successful Western blot, four requirements must be met: [do not read the bullets, go directly to paragraphs]
 Elution from the gel
 Adsorption to the membrane
 Retention during processing
 Accessibility during processing
First, the protein must elute from the gel during transfer. If it is retained in the gel, it will not be available
for analysis on the blot. Next, the protein must adsorb to the membrane during the transfer process. If
the protein is not adsorbed, it will not be available for analysis on the blot. Third, the protein must
remain adsorbed to the membrane during post transfer processing of the blot. Lastly, the adsorbed
protein must be available for antibody binding. If the protein is masked, it cannot be detected.
WESTERN BLOTTING STEPS IN A NUTSHELL
Western blotting typically consists of three main steps such as:
1. Resolution/ Separation of a complex protein sample in a polyacrylamide gel
2. Transfer of the resolved proteins onto a membrane
3. Identification of a specific protein on the membrane
But we will be breaking down these steps into a total of six procedures for better
understanding. These are based on the Western Blotting Handbooks provided by the
websites, ThermoFisher and Merck.
SPECIFIC STEPS IN WESTERN BLOTTING
1. Electrophoretic separation of proteins
Gel electrophoresis is a technique in which charged molecules, such as protein or DNA, are
separated according to physical properties as they are forced through a gel by an electrical
current. Proteins are commonly separated using polyacrylamide gel electrophoresis (PAGE) to
characterize individual proteins in a complex sample or to examine multiple proteins within a
single sample.
When combined with western blotting, PAGE is a powerful analytical tool providing information
on the mass, charge, purity or presence of a protein. Several forms of PAGE exist and can offer
different types of information about the protein(s) of interest.
Several buffering systems or gel chemistries are available for protein gel electrophoresis. Each
system provides unique advantages when resolving proteins of different molecular weights.
Separation of Complex Protein Mixtures in 1-D or 2-D Gels
One-dimensional (1-D) sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) is
commonly used to separate proteins by molecular weight prior to blotting (Figure 4).
 In some cases, non-denaturing conditions are used to separate native proteins.
 Although this method usually lacks the resolution of denaturing electrophoresis,
it may be useful when the primary antibody only recognizes non-denatured
proteins or when the protein’s biological activity must be retained.
Two-dimensional (2-D) gel electrophoresis is the technique of choice for analyzing protein
composition of cell types, tissues and fluids, and is a key technology in proteomics (Figure 5).
 Immunoblotting of 2-D gels provides information on molecular weight and
isoelectric point and can be useful to discriminate protein isoforms generated by
post-translational modifications.
 In some cases, protein phenotyping can be achieved by immunoblotting after only
a 1-D separation by isoelectrofocusing
Molecular Weight Markers
The inclusion of molecular weight (MW) standards, or markers, facilitates the estimation
of the sizes of the proteins of interest after resolution by electrophoresis.
Two types are available, unstained and pre-stained.
 Unstained MW markers usually consist of a mixture of purified native or
recombinant proteins of defined molecular weights. Visualizing their
location on a gel or membrane requires a staining step.
 Pre-stained markers
o allow monitoring of protein separation in the gel during
electrophoresis and indicate transfer efficiency in subsequent
blotting steps.
o However, they can be relatively expensive and the addition of
dyes may affect protein mobility.
o may be less accurate for molecular weight determination, as dyes
attached to the proteins may alter their ability to adsorb to the
membrane during blotting.
Polyacrylamide Concentration
Polyacrylamide concentration can be homogenous throughout the gel or a gradient. The
most common polyacrylamide concentration, 10%, is best suited for the separation of
proteins in the range of 10 – 150 kDa. If unknown proteins are being analyzed or a broader
range of separation is desired, gradient gels are recommended. For example, 4 – 12% Trisglycine gels are suitable for proteins in the range of 30 to 200 kDa, while 10 – 20% gels will
successfully separate proteins from 6 to 150 kDa. SDS-PAGE gels are usually 1.0 and 1.5
mm thick; however, for blotting and proteins transfer best out of thinner gels (= 1 mm).
Gel Running Buffers
Gel running buffers are typically composed of trisglycine or tris-tricine and may contain
0.1% detergent, usually SDS. Tris-glycine buffer systems are useful for separating a wide
range of protein molecular weights (6 – 200 kDa) and are compatible with denaturing or
non-denaturing conditions. Tris-tricine systems are best for the separation of smaller
proteins (< 10 kDa) that need to be reduced and denatured prior to loading. Both buffer
systems are compatible with protein transfer to PVDF membranes.
2. Transferring proteins to a membrane
Following electrophoresis, the protein must be transferred from the gel to a membrane.
There are a variety of methods that have been used for this process that include, but are not
limited to, diffusion transfer, capillary transfer, vacuum blotting transfer, and electroelution.
The transfer method that is most commonly used for proteins is electroelution or
electrophoretic transfer because of its speed and transfer efficiency.
Protein transfer from gel to membrane is necessary for two reasons:
 better handling capability offered by the membrane than the fragile gel during
western blot processing, and
 better target accessibility on the membrane by macromolecules like antibodies
Transfer of Proteins from Gel to Membrane
So, in simpler terms, the process of transferring proteins from a gel to a membrane while
maintaining their relative position and resolution is known as blotting. Blotting can be achieved
in three different ways:
1) Simple diffusion is accomplished by laying a membrane on top of the gel with a stack of
dry filter paper on top of the membrane, and placing a weight on top of the filter paper
to facilitate the diffusion process3. This method can be used to transfer proteins from one
gel to multiple membranes, obtaining several imprints of the same gel. The major
disadvantage of the diffusion method is that transfer is not quantitative and only transfers
25 – 50% of the proteins, compared to electroblotting.
2) Vacuum-assisted solvent flow uses the suction power of a pump to draw separated
proteins from the gel onto the membrane. Both high and low molecular weight proteins
can be transferred by this method; however, a smaller pore size membrane (0.2 µm) may
be needed for proteins with MW < 20 kDa, since they are less readily adsorbed by the 0.45
µm membrane3. Vacuum blotting of proteins out of polyacrylamide gels is uncommon
and is mostly used for nucleic acid transfer from agarose gels.
3) Electrotransfer Techniques
Electrophoretic elution, or electrotransfer17 is the most commonly used transfer method
in protein blotting. The principal advantages are the speed and completeness of transfer
compared to diffusion or vacuum blotting.
The two commonly used electrotransfer techniques are tank transfer and semi-dry
transfer. Both are based on the same principles and differ only in the mechanical devices
used to hold the gel/membrane stack and application of the electrical field. In tank
transfer, (Figure 6) the gel/membrane stack is completely immersed in a buffer reservoir
and current is applied. Typically run at constant voltage, mixing the buffer during tank
transfer typically keeps the current relatively constant. While this method is effective, tank
transfer is a slow technique that requires large volumes of buffer.
Transfer systems for western blotting
Wet transfer (tank transfer)
In wet transfer, transfer efficiencies
are better for lower molecular weight
proteins than higher molecular
weight proteins, with typical
efficiencies of 80–100% for proteins
between 14 and 116 kDa [2]. The
transfer efficiency improves with
increased transfer time. However,
with increasing time and the use of
membranes with larger pore sizes
(0.45 µm), the risk of transferring the
proteins completely through the
membrane increases (also known as
blow-through), especially for lower
molecular weight (<30 kDa) proteins.
Semi-dry transfer
Semi-dry transfer became available as the need for faster results became an issue
for researchers. For semi-dry protein transfer, the transfer sandwich is placed
horizontally between two plate electrodes in a semi-dry transfer apparatus
(Figure 5). The key to improving the speed of transfer with this method is to
maximize the current passing through the gel versus around it. To do this, the
amount of buffer used in the transfer is limited to that contained in the transfer
sandwich. Hence, it is critical that the membrane and filter paper sheets are cut
to the gel size without overhang and that the gel and filter paper are thoroughly
equilibrated in transfer buffer. Also, the use of extra-thick filter paper
(approximately 3 mm thickness) is helpful in certain semi-dry transfer devices
because these sheets can hold more transfer buffer.
Methanol may be included in the transfer buffer, but other organic solvents,
including aromatic hydrocarbons, chlorinated hydrocarbons, and acetone, should
not be added to avoid damage to the electrode plates. Fast-blotting, semi-dry
techniques use higher ionic strength transfer buffers and a high current power
supply to decrease transfer times to under 10 minutes. In rapid methods,
amperage is held constant and voltage is limited to a maximum of 25 V. Transfer
with traditional Towbin buffers can be preformed in a semi-dry apparatus either
at constant current (0.1 up to approximately 0.4 A) or voltage (10 to 25 V) for 30
to 60 minutes.
Dry transfer
Dry transfer methods use a transfer sandwich containing innovative components
that eliminate use of traditional transfer buffers. A unique gel matrix (transfer
stack) that incorporates buffer is used instead of buffer tanks or soaked filter
papers (Figure 8). The high ionic density in the gel matrix enables rapid protein
transfer. During dry blotting the copper anode does not generate oxygen as a
result of water electrolysis, unlike in wet and semi-dry techniques. This absence
of oxygen generation reduces blot distortion. Typically, transfer time is reduced
by the shortened distance between electrodes, high field strength, and high
current. As dry blotting does not require the setup time of wet or semi-dry
transfer, not only is the speed of transfer a major benefit, but the overall time
investment is improved.
3. Blocking nonspecific sites
The membrane supports used in western blotting have a high affinity for proteins. Therefore, after
the transfer of the proteins from the gel, it is important to block the remaining surface of the
membrane to prevent nonspecific binding of the detection antibodies during subsequent steps.
A variety of blocking buffers ranging from milk or normal serum to highly purified proteins have been
used to block free sites on a membrane. The blocking buffer should improve the sensitivity of the
assay by reducing background interference and improving the signal-to-noise ratio.
No single blocking agent is ideal for every experiment since each antibody-antigen pair has unique
characteristics. Empirical testing of blocking buffers is essential in optimizing a western blot
experiment. Frequently blocking buffers are made by researchers in the laboratory; however,
commercially available blocking buffers offer convenience.
4. Wash buffer formulations
Like other immunoassay procedures, western blotting consists of a series of incubations with
different immunochemical reagents separated by wash steps. Washing steps are necessary to
remove unbound reagents and reduce background, thereby increasing the signal-to-noise ratio.
Insufficient washing may result in high background, while excessive washing may result in
decreased sensitivity caused by elution of the antibody and/or antigen from the blot. As with other
steps in western blotting blot, a variety of buffers may be used.
Tris-buffered saline (TBS) and phosphate-buffered saline (PBS) are the most commonly used wash
buffers. In most cases, PBS and TBS solutions can be interchangeable. However, there are
situations on when to use one over the other. For example, TBS should be used when using
systems with alkaline phosphatase (AP)-conjugated secondary antibodies or when detecting
phosphorylated proteins with phospo-specific antibodies.
Occasionally, wash buffer formulations consist of a detergent such as 0.05% Tween 20 to aid in the
removal of nonspecifically-bound material. Depending on the specifics of the assay, the amount
of detergent in the wash buffer will vary, though typical concentrations are from 0.05 to 0.5% for
detergents like Tween 20. Another common technique is to add a 1:10 dilution of the blocking
solution to the wash buffer. Including the blocking agent with the detergent may help to minimize
background in the assay by preventing elution of the blocking protein from the membrane and/or
allowing nonspecific interactions to occur with the protein in solution rather than those
immobilized on the membrane.
It is important to note that detergents, like the protein solutions, can promote microbial growth.
While it is convenient to make pre-diluted stocks of detergents like NP-40, CHAPS, and Tween 20,
fungi can grow in these solutions, which can lead to high background noise. In addition, detergents
can contain significant amounts of peroxides which will cause background signal when using
horseradish peroxidase substrates. Therefore, it is important to use high-purity detergents.
5. Primary and secondary antibodies
Western blotting is typically performed by probing the blocked membrane with a primary antibody
that recognizes a specific protein or epitope on a group of proteins (e.g., SH2 domain or
phosphorylated tyrosine). The choice of a primary antibody for a western blot will depend on the
antigen to be detected and what antibodies are available to that antigen. It is also important to
note that not all primary antibodies are suitable for western blotting and the application should
be verified, if possible, before purchasing a new primary antibody.
In general, the primary antibody that recognizes the target protein in a western blot is not directly
detectable. Therefore, tagged secondary antibodies are used as the means of ultimately detecting
the target antigen (indirect detection). A wide variety of labeled secondary antibodies can be used
for western blot detection. The choice of secondary antibody depends on either the species of
animal in which the primary antibody was raised (the host species) or any tag linked to the primary
antibody (e.g., biotin, histidine (His), hemagglutinin (HA), etc.) For example, if the primary
antibody is an unmodified mouse monoclonal antibody, then the secondary antibody must be an
anti-mouse IgG secondary (or non-IgG) antibody obtained from a non-mouse host.
Antibodies for western blotting are typically used as dilute solutions, and manufacturers may
recommend using ranges from a 1/100–1/500,000 dilution from a 1 mg/mL stock solution.
However, the optimal dilution of a given antibody with a particular detection system must be
determined experimentally. More sensitive detection systems require less antibody than lower
sensitivity systems and can result in substantial savings on antibody costs and allow a limited
supply of antibody to be stretched out over more experiments. Using lower amounts of antibody
can also have the added benefit of reduced background because the limited amount of antibody
shows increased specificity for the target with the highest affinity.
Antibody dilutions are typically made in the wash buffer. The presence of detergent and a small
amount of the blocking agent in the antibody diluent often helps to minimize background, thereby
increasing the signal-to-noise ratio. Conversely, adding too much blocking agent or detergent to
the antibody dilution solution can prevent efficient binding of the antibody to the antigen, causing
reduced signal as well as reduced background.
6. Detection methods
While there are many different tags that can be conjugated to a secondary or primary antibody,
the detection method used will limit the choice of what can be used in a western blotting assay.
Radioisotopes were used extensively in the past, but they are expensive, have a short shelf-life,
offer no improvement in signal-to-noise ratio and require special handling and disposal.
Alternative labels are enzymes and fluorophores.
Enzymatic labels are most commonly used for western blotting and, although they require extra
steps, can be extremely sensitive when optimized with an appropriate substrate. Horseradish
peroxidase (HRP), and to a lesser extent, alkaline phosphatase (AP) are the two enzymes used
most extensively as labels for protein detection. An array of chromogenic, fluorogenic, and
chemiluminescent substrates are available for use with either enzyme. Alkaline phosphatase offers
a distinct advantage over other enzymes in that its reaction rate remains linear, improving
sensitivity by simply allowing a reaction to proceed for a longer time period. Unfortunately, the
increased reaction time often leads to high background signal resulting in low signal-to-noise
ratios. Horseradish peroxidase–conjugated antibodies are considered superior to antibody-AP
conjugates with respect to the specific activities of both the enzyme and antibody due the smaller
size of HRP enzyme and compatibility with conjugation reactions. In addition, the high activity
rate, good stability, low cost, and wide availability of substrates make HRP the enzyme of choice
for most applications.
Enzyme-conjugated antibodies offer the most flexibility in detection and documentation methods
for western blotting because of the variety of substrates available. The simplest
detection/documentation system is to use chromogenic substrates. While not as sensitive as other
substrates, chromogenic substrates allow direct visualization of signal development.
Unfortunately, chromogenic substrates tend to fade as the blot dries or during storage, making
the blot itself an unreliable means of documentation. However, it is fairly straightforward to either
photocopy or directly scan the blot in order to make a permanent replica of chromogenic western
blot results.
Chemiluminescent blotting substrates differ from other substrates in that the signal is a transient
product of the enzyme-substrate reaction and persists only as long as the reaction is occurring. If
either the substrate is used up or the enzyme loses activity, then the reaction will cease and signal
will be lost. However, in well-optimized assays using proper antibody dilutions and sufficient
substrate, the reaction can produce stable output of light for 1 to 24 hours depending on the
substrate, allowing consistent and sensitive detection that may be documented with X-ray film or
digital imaging equipment. While X-ray film can be used to obtain semi-quantitative data, digital
imaging is more sensitive because of the broad dynamic range of detection, allowing researchers
to obtain quantitative data from western blots.
The use of fluorophore-conjugated antibodies requires fewer steps because there is no substrate
development step in the assay. While the protocol is shorter, this method requires special
equipment in order to detect and document the fluorescent signal due to the need for an
excitation light source. Recent advances in digital imaging and the development of newer
generation fluorophores such as infrared, near-infrared, and quantum dots has increased the
sensitivity and popularity of using fluorescent probes for western blotting and other
immunoassays. Although the equipment and fluorophore-conjugated antibodies can be quite
expensive, this method has the added advantage of multiplex compatibility (using more than one
fluorophore in the same experiment). In addition, chemical waste is further reduced compared to
other blotting procedures.
Western Blot Detection Considerations
Western blots detect specific protein from cells or tissues
in a convenient, flexible format for rapid evaluation.
The western blot format can also be quantitative and offer
a high degree of sensitivity. With a variety of detection
techniques, including chemiluminescent, fluorescent, or
chromogenic to choose from, you can select a technology
to match your experimental requirements and the
instruments you have available. We discuss below a few
key factors to consider before performing western blotting.
Signal-to-noise ratio
Signal-to-noise ratio compares the level of desired or
relevant signal to the level of background noise or irrelevant
signal; the higher the ratio, the better the result. In western
blotting, the signal is the density of the specific probed
protein band of interest; the noise is the density of the
background. In western blotting applications, optimization
of the signal-to-noise ratio is often more important than
increasing the sensitivity of the system. The sensitivity of
the system is irrelevant if the signal cannot be adequately
distinguished from the noise. For information on western
blot optimization methods, see page 80.
Direct vs. indirect detection
The antibody that recognizes a target protein is called the
primary antibody. If this antibody is labeled with a tag for
visualization purposes (typically an enzyme or fluorophore),
direct detection of the target is possible. Typically, the
primary antibody is not labeled for direct detection.
Instead a secondary antibody that has been labeled with
a detectable tag is used to probe for the primary antibody,
which is bound to the target. Thus, the target is detected
indirectly. Indirect detection with secondary antibodies
requires more steps than direct detection, but it can also
offer significant advantages over using primary antibodies
that are directly labeled (Figure 13).
Indirect methods can offer increased sensitivity through
the signal amplification that occurs as multiple secondary
antibody molecules bind to a single primary antibody. In
addition, a given secondary antibody will recognize most
primary antibodies of the same isotype and target species,
making it a more versatile reagent than individually labeled
primary antibodies.
Several variants of these probing and detection strategies
exist. However, each variant depends on a specific
probe (e.g., a primary antibody) whose presence is linked
directly or indirectly to some sort of measurable tag.
In this handbook, most methods discussed use indirect
detection, as this has emerged as the most popular
detection strategy.
Manual vs. automated western blot processing
Traditionally, probing a western blot prior to data
visualization involved a series of manual steps, many of
which were individually short but collectively required
significant hands-on time. Today, instruments are available
to automate some of these tasks, tremendously decreasing
hands-on time.
Manual and automated procedures share three essential
steps: blocking the membrane, probing with primary
and secondary antibodies, and washing the membrane