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.