Western Blot

This is going to be a long and technical one sorry. 

This month I want to talk about an experiment I've done pretty much every day for the last three months (plus continuously over the last 2.5 years) - Western Blot. 

Western blots are used to visualise and identify individual proteins. In this technique you have a solution (called a lysate) filled with the majority of proteins from cells. This technique separates these proteins out by size and allows you to stain for specific proteins you want (more on this later). This is the standard way to visualise proteins but it is very subjective and is tricky to perfect (I hope you will appreciate why after reading this). 

The proteins are visualised by staining them with antibodies specific to that protein and then using a chemical reaction to produce light which can be picked up by a machine. The resulting image is of black bands. The intensity of the black band determines the approximate level of protein present. 

There are a few steps to Western Blotting which I will go through. The whole technique can take 1-3 days depending on what proteins you are looking for and what you choose to do in each step. 

Steps to Western Blotting:

1. Gel preparation, protein preparation and buffer preparation
2. Loading gel and running SDS-PAGE gel electrophoresis 
3. Transferring proteins onto membrane
4. Blocking non-specific binding
5. Antibody staining of proteins
   1. Primary antibody incubation and washing
   2. Secondary antibody incubation and washing
6. Developing 

A little bit of terminology before we begin:

• Blot: a blot is what we call the membrane once protein has been transferred to it (more later). A blot can be the whole gel you ran in electrophoresis or part of that gel. 
• 1°Ab: primary antibody 
• 2°Ab: secondary antibody 
• Loading control: to make sure that you have loaded the same amount of protein in each well and to ensure that any changes in protein abundance in response to say a drug is real you also stain for a loading control. This is typically an abundant protein found in every cell. Usually beta actin or tubulin are used. 

STEP 1: Preparation

Reagents and Buffers
Before you begin a western you need to make sure you have all your reagents prepared. I will NOT be going through this, all I will say is you need reagents for

• making your gel
• making up your protein that you will load onto the gel (loading buffer)
• the buffer the SDS-PAGE gel electrophoresis will run in (running buffer)
• the TBST you will use to wash your blots (wash buffer)​

Protein Preparation:

When you extract your protein from your cells you end up with a protein lysate. This contains the majority of the proteins in your cells. You don't know what the concentration of the total protein is but you are sure that each sample doesn't contain the same concentration. This is due to how well the extraction went in each individual sample but also how many cells there is to extract protein from. There are a few ways of finding out the concentration of your protein which I won't go through (I use the Bradford Assay from BioRad). 

What you do need to know is that for each sample you need to make up a solution of protein, water and loading buffer to load onto your gel. You can use as much or as little protein as you want, I typically use 10-15ug per well. I load 20ul per well so I need:

• Protein - e.g. 5ul
• Water - e.g. 10ul
• Loading buffer - ALWAYS 5ul per well 

​There are many types of loading buffer, I use Laemmli buffer.
The loading buffer has a couple of functions. You are using an electric current to make the proteins travel through the gel. Proteins are all different charges so the buffer makes sure all the proteins have the same charge (negative) - this is done by a chemical called SDS. Proteins are globular structures. These don't run though a gel very well so you need to unravel the proteins into single strands (called denaturation).  The buffer will start this process - a chemical called β-mercaptoethanol breaks disulphide bonds. The loading buffer also has a dye in it. This is called the "loading front" which runs faster than your proteins. This allows you to track how far your proteins have travelled through the gel (image later). ​

Once you add the protein, water and loading buffer together you boil the protein at 95-99°C for 5 minutes. This completes the denaturation of the proteins. The samples are kept on ice until needed to prevent further degradation of the proteins. 

A quick note - usually the protein samples you have prepared will be blue once you add the loading buffer and boil it. This is because the dye in the loading buffer is blue. However, the protein I will show in this post will be yellow. This is because I extracted the proteins using acid and the acid in the lysate turns the buffer yellow. Once the gel starts running the dye front goes back to blue.

Gel Preparation:
​
​The first step of western blotting is to separate out your proteins (called resolving). This is done using a gel which is sort of the consistency of jelly you would eat except you do not eat it...

Image of reagents used to make stacking and resolving gels for electrophoresis

Gels are made up of:

• 30% Acryl-bisacrylamide mix - polymerizes to form a matrix, which allows proteins to pass through
• 1.5M Tris (pH 8.8 or pH 6.8) – allows a negative charge to be applied to proteins
• 20% SDS - denatures secondary and tertiary protein structures and adds a negative charge. 
• 10% ammonium persulfate – an oxidising agent often used with TEMED to catalyse polymerisation of acrylamide (Added just before setting gel)
• TEMED - stabilises free radicals produced by APS and improves polymerisation (Added just before setting gel)

The acrylamide will set into a gel which has pores in it. The protein will travel through these pores. The percentage of acrylamide used determines how big these pores are. Large proteins run slowly through a gel whereas smaller proteins will run very quickly through the gel. A lower percentage gel (e.g. 7%) will have large pores, allowing larger proteins to run faster through the gel. However you are likely to lose all the smallest proteins. A higher percentage gel (e.g. 12.5%) will have very small pores which slow down the small proteins, allowing you to visualise them easier. However all your largest proteins might not run as they can't past through the pores. 

There are two types of gels used in gel electrophoresis - a stacking gel and a resolving gel. They're pretty much made up of the same ingredients (a stacking gel is slightly more acidic pH 6.8). 

• The stacking gel: is usually a very low percentage gel (4-5%), meaning it has very big pores. This is where the comb is added to form wells which allow you to load your protein into the gel. The stacking gel allows all the protein to line up, like sprinters on a start line, so the proteins run smoothly and evenly through the gel. 
• The resolving gel: this is where your proteins separate by size (resolving). The percentage of this gel depends on the proteins you are looking at. 

Image of gel set up with comb

Gels are made in a cassette (shown on the left). A cassette is made up of two pieces of glass sandwiched together by the green clamp. One piece of glass (the backing plate) has a space in the middle. The cassette is placed in the stand which creates a seal at the bottom to prevent the liquid from leaking out

Steps to making a gel:

1. Make up your cassette in the stand
2. Add the resolving gel FIRST and allow to set. Typically you put a little isopropanol on the top (which won't mix with the resolving gel) to prevent evaporation while the gel is setting
3. Remove the isopropanol and wash with water 
4. Add the stacking gel 
5. Place in a comb into the stacking gel. This creates the wells for loading protein. The number of wells depends on the comb you use - typically 10-15 wells. 
6. Allow to set

I have made up a 12.5% gel for this example to run histone proteins which are approx 15-17kDa in size. I will also stain for a loading control, beta actin, which is 45kDa in size. 

STEP 2: Loading and running SDS-PAGE Gel Electrophoresis

The next step is to load your prepared protein into the wells in your stacking gel and running an electric current through the gel to allow the proteins to travel from negative to positive. 
​
The gel(s) are clamped into an electrode chamber and placed in a running tank. The centre is filled with running buffer. The running buffer allows an efficient flow of electrical current to pass through the gels. It also prevents the gels from drying out which you do not want. 

Image of electrophoresis tank with gels.

Image of capillary tip

The proteins samples are loaded into each well using a capillary tip (shown above). This allows for more accurate pipetting. As I've said before each well will have 20ul sample with 10ug protein per well.

To load the well you first take the comb out carefully, place the pipette tip with your sample into the well and very slowly and carefully release the sample into the well. The wells are pretty hard to see so you have to get very good at guessing where the next well starts, especially when you're adding a yellow liquid into a clear liquid. 

Videos of how a gel is loaded

Image of gel as electrophoresis begins. Samples are still dispersed in wells and have no run through the stacking.

At the start and end of each row of wells is a black liquid. This is called a ladder - the ladder has coloured bands which correspond to approximate protein sizes (you will see this more clearly later). This allows you to cut your membranes to optimise protein visualisation from a single gel (you can't look for more than one protein at a time) and also allows you to semi-accurately estimate the size of your protein once developed. 

Once the samples are loaded they are run through the gel in two steps:

1. the samples run through the stacking to all line up - this is done at 80V for about 10-15 minutes
2. the samples run through the resolving gel - this is done at 100-120V (i usually do 110V) for 1-2 hours. 

Image of protein samples after running through stacking gel. The ladder starts to separate out (resolve) and the dye front returns to blue.

STEP 3: Transferring proteins from gel to PVDF membrane

Once the gel has run enough you can stop it. This depends on how long you want to wait for and what proteins you're looking for. My proteins are very small (17kDa) so I can stop my gel once I see the 17kDa ladder band is far enough from the loading front. 

The next step is to transfer the proteins from the gel onto a membrane. You cannot stain for your proteins on the gel as it is far to delicate and not permeable - the protein only runs through the gel because we created the wells. The membrane is more permeable and allows you to stain for proteins. 

There are a couple of (messy and time consuming) ways to transfer proteins but I prefer to use the iBlot dry technique. This basically transfers the proteins using a machine in 5 minutes (compared to 1-2 hours). The transfer stack consists of:

1. bottom part: which has the membrane (in this case a PVDF membrane)
2. middle part: the gels, covered in a piece of damp filter paper
3. top part: allows electricity to be conducted
4. sponge

Again an electric current is used to push the proteins through the gel into the membrane. However the voltage is very low. As we know, small proteins move very quickly. The low voltage prevents the small proteins from passing straight through the membrane. 

So how does it work:

1. You remove the gel cassette(s) from the tank
2. You gently open the cassette. You cut off the stacking gel and the loading dye front. The gel is placed in water until ready to transfer to prevent it from drying out
3. The bottom stack is opened and the gel(s) are placed on top
4. A piece of filter paper is quickly wetted and placed on top of the gel(s)
5. A roller is used to gently remove any air bubbles and excess water
6. The top stack is placed on the filter paper (metal side up)
7. The whole completed stack is placed in the iBlot machine, with a sponge placed in the lid
8. The lid is closed firmly, ensuring there is a complete current
9. The transfer is run at 20V for 5 minutes

Image of how you set up before transfer

Image of bottom transfer stack with gels on top of the membrane

The iBlot transfer machine with the stack and sponge in place

​Videos of how the transfer stack is prepared

After the transfer is complete you need to be VERY fast. You have to quickly remove the stack from the machine, get the membrane out of the stack and place it in TBST. The membrane CANNOT dry out or else all is lost. You can check your transfer at this stage using Ponceau red, which is a red dye that shows up all the wells and bands of proteins but I did not do that today. 

STEP 4: Blocking

The next step is to block any unspecific binding. The antibodies are usually polyclonal - meaning they can bind to other proteins that aren't your protein of interest. To prevent this and also to get a nice clean image at the end, you block unspecific binding. 

You can block using either of these two reagents:

1. 5% BSA/TBST/NaN3
2. 5% Milk/TBST

The reagent you are using depends on the antibodies and proteins you are looking for. I block in 5% BSA/TBST/NaN3.

Before I block I cut my membrane so that I have the two individual gels again. You don't have to do this but I like to. At this point I just cut the excess off using the ladders as a guide. I then mark with a pen on the ladder what gel it is. 

The membranes are blocked for 1 hour at room temperature on a rocker. 

This is now when we can call out membrane a blot. 

After blocking you can cut your blots to stain for multiple proteins. In this case I cut at 36kDa band (blue band above the bottom red band on the ladder) to stain for my histone proteins and also my beta actin loading control. 

Image of membranes after being cut

Image of membranes blocking in 5% BSA/TBST/NaN3 for 1 hour at room temperature on a rocker

STEP 5: Antibody Staining of Proteins

This technique as I said uses antibodies which bind to specific proteins and you can them visualise these antibodies using a chemical reaction. 

​ The figure below gives a rough idea of how the technique works. 

1. a primary antibody semi-specific to the protein of interest binds to a particular part of the protein.
2. a second antibody with a HRP (horse radish peroxidase) attached to it binds to the first antibody. 
3. A peroxide solution is added which reacts with the HRP. This chemical reaction produces light which is detected using chemiluminescence. NOTE: the luminol enhancer solution is added to increase the efficiency of the chemical reaction making it easier to see. 

The antibodies are created in different animal models (please don't ask how cause I have no idea). So if your primary antibody is developed in a rabbit then the secondary antibody will be goat anti-rabbit (i.e an antibody developed in a goat to bind specifically to rabbit-developed antibodies). There are some human antibodies but they're incredibly expensive. 

The primary antibodies are made up in the buffer recommended by the manufacturer usually 5% BSA/TBST/NaN3. The recommended concentration is usually given by the manufacturer - typically 1:1000 or 1:2000 (i.e. 1ul antibody in 1ml solvent).  
The secondary antibodies are made up in usually either 5% BSA/TBST or 5% Milk/TBST. The BSA doesn't contain NaN3 in the secondary because sodium azide (NaN3) inhibits the secondary antibody. The concentration of the secondary depends on how abundant the protein is and how long you develop it for. For example beta actin, which makes up part of the cytoskeleton of your cells, is an incredibly abundant protein so the secondary concentration is 1:50,000 BUT another protein I look at p-AMPK is not so abundant and the secondary concentration could be as low as 1:1000. 

Diagram of antibody staining of proteins

Part 1: Primary antibody incubation and washing

If you protein is not hugely abundant or you don't want to do the whole process in one day, you can incubate your blot(s) overnight in your primary antibody at 4°C on a roller. 
Once the blot(s) have been incubated in the primary antibody you need to wash the blot(s) at least 3x10 minutes in TBST. This removes all the excess primary antibody. If you put the blot(s) immediately into the secondary your final image will come out black. The washes are done on a rocker at room temperature.

NOTE: for beta actin, you do a 1 hour incubation at room temperature

Part 2: Secondary antibody incubation and washing
​
​For the secondary antibody you can do this at room temperature for 30 minutes - 1 hour, again on a roller.
I make up most of my secondaries in 5% Milk/TBST, except p-AMPK. The casein in the milk removes phosphorylation from proteins so I would lose my p from the p-AMPK if I did the secondary in milk. So for p-AMPK and all phosphorylation proteins I use 5% BSA/TBST. 

Again after incubation in the secondary, the blot(s) need to be washed at least 6x5 minutes to remove excess secondary antibody.

The washing part of the procedure is honestly the most time consuming and tedious. It's a lot of back and forth to and from my office desk to the rocker. I usually try to do something else in the lab to not waste the time but sometimes it can't be avoided. 

STEP 6: Developing

The final step (thank god) is to develop your blot. This step adds the peroxide and luminol enhancer solutions (collectively known as ECL solutions) to the membranes. You then take the membranes to a chemiluminescence machine which will detect the light emitted and give you an image in return. 

Steps:

1. an equal amount of peroxide solution and luminol enhancer solution are added together. 
2. The blot(s) are dried slightly on tissue and placed in the ECL solutions
3. The blot(s) are incubated for 1 minute in the ECL solutions 
4. The Blot(s) are transferred to a clear plastic sheet and covered with another clear plastic sheet
5. Excess bubbles are removed
6. The blot(s) are placed in the chemiluminescence machine and developed for maximum 15 minutes. This can be done in increments (e.g. 1 minute increments for 10 minutes) ​

Image of the ECL technique

Image of blot ready for chemiluminescence

Image of the chemiluminescence machine

You end up with a picture which has black bands. As I said at the start the intensity of the bands indicates how much protein is present - very dark means a lot, very light means very little and nothing means nothing. ​

Image of a beta actin western blot for MCF10A (lane 1-3) and MCF12A (lane 4-6).

The image above shows the loading control (beta actin) for a blot I have done for two cell lines, MCF10A (lane 1-3) and MCF12A (lane 4-6). The bands are very dark indicating that the protein is very abundant. What's important is the bands are almost the exact same size and intensity as each other (different in the two cell lines) indicating that my loading was very even (wahey). 

Once the blot is developed you can keep it or you can throw it away. If you keep it and want to stain for another protein or the same protein but with a different secondary concentration you need to strip the blot (i.e. remove all antibodies bound to proteins), re-block using the same blocking buffer and re-incubate with the primary and secondary antibodies. A blot should only really be stripped twice. This will add another day onto your experiment (at least). 

And that is it. That is roughly how a western blot works. 

The reason why western blots are THE most hated laboratory experiment is because they rarely go the way you want them to, there is always a difference each time you do it and it takes FOREVER. 

If a western goes wrong most of the time you have no idea why. It can be anything from one specific buffer which was made up wrong to the wrong gel percentage, the wrong transfer time or transfer voltage etc. Basically everything you did could have an issue associated with it. Trust me I did troubleshooting for TWO MONTHS on one antibody. It eats away at your soul. 

Western blotting everyone.