Welcome to the Novus Visual Protocol Series.
In this video we will learn how to perform all phases of a Western Blot using the
most common methods for this assay.
Before we can start preparing the blot we must first prepare our sample lysate.
In this example we will prepare a protein lysate from cultured cells.
Here
we wash the cells twice with ice cold PBS
and enough lysis buffer to cover the cells.
The choice of lysis buffer depends largely on your
choice of protein of interest.
We scrape the cells and transfer the cell solution on a centrifuge tube
placed on ice.
In order to solubilize membrane bound proteins, we will require stronger
extraction detergents compared to isolated cytoplasmic proteins.
In this example
we are using a standard RIPA buffer,
which is a common buffer for obtaining maximum protein yield. While extracting
proteins from all cellular localizations,
it is very important to include protease inhibitors in your lysis buffer which will
prevent degradation of your sample. Always use freshly prepared protease
inhibitors, keep samples on ice and work quickly.
We lice the cells by pipetting up and down
followed by incubation on ice for 30 minutes.
Then centrifuge the cells into a pellet.
Discard the pellet and collect the supernatant. This is your lysate.
Determine the total concentration of your protein lysate by
testing a small portion of your lysate
with a commercially available protein quantitation assay
such as the BCA.
This will assist you in loading equal amounts of protein into your gel.
Western Blots are typically preformed under reduced and denatured
conditions. These conditions will allow proteins to be separate by their
molecular weight
rather than their native conformational shape or charge.
To reduce and denature samples,
dilute each in a loading buffer such as the traditional laemmli buffer.
This buffer contains beta-mercaptoethanol, or DTT, to reduce disulfide ridges
between cysteines,
SDS to assist denaturing a net negative protein,
glycerol to allow the samples to sink into each well,
bromophenol blue to visualize the lysate and an iconic buffer.
Votex each sample at 95 degrees Celsius for five minutes to
completely denature the proteins. You are now ready to load your samples
into an SDS page gel.
For this next step we will separate the individual proteins in our sample
lysate
based upon their molecular weight
using a positive electrode to attract a negatively charged protein.
To do this
we load our previously prepared protein samples into a commercially available
polyacrylamide gel.
Gels are available in fixed percentages or gradients of acrylamide.
The higher the acrylamide the smaller the proportion of gel
percentage.
Therefore higher percentage gels are better for low weight proteins, low percentage
gels are better for large weight proteins and gradient gels can be used for
proteins of all sizes
due to their varying range in pore size.
Prepare your gel by inserting it into the electrophoresis apparatus and
filling with running buffer that is appropriate for your gel chemistry.
Rinse the wells of the gel with running buffer and add buffer to the
chambers.
Load your samples into the wells.
If you are unsure of the amount to load,
10-30 micrograms of total protein is a suggested starting point
as well as the entire amount of sample loaded.
You will also need to reserve at least one well for prestained molecular weight
ladder.
The ladder will allow you to monitor protein separation during
electrophoresis and subsequently verify protein weight in your sample during
later analysis.
Close the electrophoresis unit and connect it to a power supply. Most units
typically run 45-60 minutes at 200 volts
or until the loading buffer reaches the bottom of the gel.
During this time the negatively charged proteins in each sample will migrate toward
the positively charged electrode making their way through the polyacrylamide
gel matrix.
In this next step, we will transfer our separate proteins out of the gel
and into a solid membrane or blot.
This is based upon the same principal as the previous step
in which an electric field is charged to remove the negative proteins
towards a positive electrode.
Transfer can occur under wet or semi-dry conditions.
Here we will demonstrate the traditional wet transfer method. Start by removing
the gel from its cassette
cutting the top portion containing the wells.
Notch the top left corner to indicated gel orientation.
Float the gel in transfer buffer while preparing the transfer sandwich.
To make the transfer sandwich
you will need a cassette,
sponge, filter paper
the gel
and your choice of either PVDF or nitrocellulous membrane.
PVDF must first be activated by soaking the membrane in ethanol for
two minutes. But other than this the PVDF or choice of nitrocellulous
membrane is a personal preference.
Notch the top left corner to indicate blot orientation
and incubate membranes in transfer buffer for 10 minutes.
Create a stack by placing the following components
from the black negative cathode to red positive anode:
sponge,
filter paper,
membrane,
(Be careful not to touch the gel or membrane with your bare hands and use
clean tweezers or spatula instead.
Touching the membrane during any phase can contaminate the blot and lead to
excessive background signal. ),
filter paper
and sponge.
Use a clean roller with each layer to gently roll out any bubbles that may be
present
since bubbles will inhibit efficient protein transfer.
Lock the cassette and place it in the apparatus containing cold
transfer buffer
ensuring that the cassette is properly positioned from negative to positive.
In order to prevent heat buildup, it is beneficial to transfer with a cold
pack in the apparatus
apparatus or in a cold room with the spinner bar placed at the bottom of the chamber.
Close the chamber and connect to a power supply.
Perform the transfer according to the manufacturer's instructions which is
normally 100 volts for thirty to one hundred and twenty minutes.
After electrotransfer of our proteins to a membrane, we will
now block the blot,
apply a primary antibody specific for our protein of interest and then a secondary
antibody which will recognize the primary antibody.
Start by removing the membrane from the cassette and rinsing three times in water.
As an optional step, we can verify the proteins were transferred successfully
by staining the membrane with ponceau red.
Incubate the membrane in ponceau for five minutes and wash with water until
the bands are clear.
After verification
the blot can then be de-stained by continuing to wash with water
or TBS twine
until the dye is completely removed.
We need to block all areas of the blot which do not contain protein.
This will prevent non-specific binding of the antibody and reduce overall
background signal.
Common blocking buffers include 5% non-fat dry milk for the assay
in a TBS-Tween solution.
However do not use a milk solution when probing with phosphor-specific antibodies
as it can cause high background from its endogenous phosphoprotein, casein
Incubate the membrane with blocking solution for one hour at room
temperature
under slight agitation.
Decant the blocking solution and wash with TBS twine for five minutes.
We are now ready to add our antibody. Dilute the primary antibody in a
blocking buffer at the concentration recommended on the datasheet.
Incubate overnight at 4 degrees Celsius with gentle shaking.
A recommended optional step is to also use a positive control antibody
which allows the user to verify equal amounts of total protein were loaded into
each well and aides in troubleshooting by removing any
uncertainties with the Western Blot procedure.
The next day,
decant off the primary antibody and wash the membrane with large volumes
of TBS twine and vigorous agitation
five times for five minutes each.
These stringent washes are extremely important for removing non-specific
background signals.
After washing, dilute the secondary antibody in blocking solution and incubate
the membrane for one hour at room temperature at the concentration
recommended on the datasheet.
In our example the secondary is also conjugated to HRP for later
detection.
Decant membrane and wash secondary with large volumes of TBS twine with
vigorous agitation five times for five minutes each.
You are now ready for the detection phase.
In this final phase, we will demonstrate signal development using the most common,
most sensitive and most inexpensive detection method the electrochemiluminescence
(or ECL) reaction.
This method utilizes the HRP enzyme,
which was conjugated to the secondary to catalyze the ECL reaction and produce
light.
A light is then gathered onto x-ray film and developed
or digitized with the aid of a specialized camera sensitive enough
for this application.
We start by mixing equal parts ECL reagents in a one-to-one ratio
according to the manufacturer's instructions.
We will incubate the membrane for 3-5 minutes without agitation.
After incubation, decant ECL mixture and use a laboratory wipe to wipe off excess
solution from the corner of the membrane.
Place the membrane in a clear plastic wrap such as a sheet protector to prevent
drying. Avoid letting the membrane completely dry out.
We can now use a roller to push out any bubbles or any excess solution.
Immediately develop the membrane.
Both film and camera systems allow you to manually adjust the exposure time
in order to ensure a picture perfect Western Blot.
Relative band densities can now be quantified with commercially available
software.
. Proper molecular weight can also be verified by comparing band sizes to the
molecular weight ladder.