Honest Protocols: Antibodies, visualising, and stripping

 We finally made to the end of the western blot honest protocols series!  

At this stage, our membrane has been left to block in the cold room overnight and now we are going to start working towards the visualisation stage - where we will find out two things:  the protein you wanted in the first place was in the lysate, or the western blot didn't work.

So, to see if the protein is present, we are going to use antibodies. Antibodies are the reason your western blot looks like it does and produces those bands. Each of those bands is where an antibody has bound to your target protein. 

There’re two types: the primary and the secondary.

The point of the primary is to bind to the target protein - or the protein that you want to know is within your lysate. Most of the primary antibodies are monoclonal -meaning they will only bind to one protein (in theory) meaning that you need different primary antibodies for every protein you want to detect. 

The secondary binds to the primary antibody and is there to amplify the connection between the primary and the protein of interest. Whilst the primary antibody finds the protein, the secondary antibody makes it visible. The secondary antibody isn't specific, and like the primary antibody is usually produced in a rabbit or goat. Weird but there we go. If you use a goat primary antibody, you also have to use a goat secondary. Same goes for rabbit. 

Handy diagram on how primary and secondary

antibodies work. 

So, what you need to do is apply the primary antibody to the membrane, leave it for a while and then add the secondary antibody. In between this, you want to be washing off the excess antibody to stop any background noise. You also want to be washing off the blocking solution. 

So: 

1) Pour away the blocking solution and leave your membrane floating in PBS-T* on the shaker in the lab (not the one in the cold room). 

*PBS-T is phosphate buffered saline with a small amount of Tween-20, a detergent added. It keeps the pH stable, and the Tween helps prevent the antibody from binding to anything it shouldn't. You can also use TBS- which is Tris buffered saline. Both work more or less the same.  

2) Leave it for five minutes.

3) Pour off the PBS-T and add more PBS-T. 

4) Repeat steps 2-3 about three times. Lose track of how many times you have actually done it. 

5) Check to see whether you still have some antibody solution left over. Much to your disappointment but not to your pressure, you don't. 

 6) Sigh when you realise that this means you need to make 5% BSA-PBS-T solution.

   *BSA-PBS-T is BSA mixed with PBS-T.  Typically, it's about 2.5g with 50ml of PBS-T. The best way to do it is weigh out 2.5g of BSA, add it to a tube and then top it up into 50ml.

7) Pause briefly to wipe up the BSA you split all over the balance.

8) Spend the next 10 minutes frantically shaking the tube up and down in an attempt to dissolve the BSA. 

9)Have a rare moment of forward thinking and remember that you may need more of the 5% BSA-PBS-T. So, make up another tube and stuff in the fridge for later.

10) Retrieve your vial of primary antibody from the -20 freezer. 

11) Start thinking about what concentration of antibody you will actually need- and how much of the antibody solution you will need. Decide on 15 ml of BSA-PBS-T and add 15 ul of antibody*

  *Typically, you use a concentration of 1:1000, for the primary antibody and see what happens. You can try and increase the concentration later. Some antibodies work with a lower concentration, or some need a stronger. concentration. There's no way to know until you try it. 

12) Pour away the PBS-T on your membrane and add about 10 ml of the antibody solution onto the membrane. Like with blocking, check the membrane is moving. 

13) Incubate with primary antibody overnight on shaker in cold room*

 *Or one hour on the shaker at room temperature. Depends on how lazy you are being. But it has to be at least one hour if doing it at room temperature.

14) Wash the membrane with PBS-T five times. 

 15) Retrieve your secondary antibody from the 4 degrees freezer and make up your secondary antibody solution* with 5% BSA-PBS-T.

      *The secondary antibodies are stronger so use a concentration of about 1:3000. Which means about 5ul for 15 ml. And remember to check whether your primary was a goat or a rabbit. Your secondary has to be the same. 

16) Remove antibody from membrane*. 

   * Just pour it back into a conveniently labelled test tube (for the love of God, make sure the test tube is labelled). The antibody can be re-used but no more than 5 times at the absolute most. 

17) Pour about 10 ml of secondary antibody on to the membrane and leave it to it for 1 hr on shaker at room temp*.

* No matter how lazy you are feeling, the secondary cannot be left on overnight. It causes background noise.

18) Wash three times in PBS-T, like you did with the primary.

19) Take a deep breath and brace yourself for your fate. After all this work and hassle, you get the fun of visualising it and seeing if you all your hard work was for nothing or not.

20) Sadly wander over to the visualiser to book a slot. 

21) Unfortunately (or perhaps fortunately), it's not available right now.

22) Book a slot for about an hour from now and leave your membrane sulking/soaking in PBS-T*,

*You could leave the membrane soaking in PBS-T for a week and it would be fine (That is not scientific advice).

23) Come back after an hour and grab your ECL from the cold room.

   *ECL is the detection reagent you will use to see your protein bands. Your secondary antibody usually has something called HRP attached to it. This reacts with the ECL and gives off a small amount of light. It's not visible to the human eye- so you need the gel visualiser to see it. This light is what forms a band. Which means in theory, you only get the bands where the proteins are.  You can then use the ladder to determine how big the bands are. This is how you can confirm that the band is the protein you actually wanted. 

How ECL works on the primary antibody and 

secondary antibody. 

24) Just before* you are about to use it, make it up using a 1:1 ratio of reagent A to B. 

*ECL degrades quickly so you don't want it sitting around for long. Depends on the membranes but I usually made it up with 5ml reagent 5 A and 5ml reagent B. I'm not making that up - they are actually called A and B. 

25) Forget all you know about ratios.

26) Eventually figure it out. 

27) Nervously approach the visualiser. Tentatively turn it on and try to remember how to use it*.

*Every visualiser is different but there's usually instructions nearby. 

28)ECL didn't show anything, so you try with ECL plus*

   *ECL Plus is basically a more sensitive version of ECL. Sometimes, a protein might be on the membrane but at a really low amount so you need to use something more sensitive.  For ECL plus, you use 1:40 ratio of reagent A to B. 

29)That also doesn't work so you decide to try West Femto.

*West Femto: Even more sensitive. If you are at this point, there's not that much hope but there is still a possibility!!  1:1 ratio of reagent A to B

30) Pray. Pray. Pray.

31) Pray again to the western blot gods. They do exist and they are watching over you.

32) Let's pretend this hypothetical western worked and you actually got the bands you wanted. Happy days! 

What a successful Western should look like.

Either side is the ladder whilst the thicker black band

 about a third down is the protein of interest. Some background bands 

can also be seen but they aren't getting in the way.

33) Soak membrane in PBS-T for three times, 5 minutes each. 

34) Find the stripping buffer*

* You want to remove the secondary antibodies from the membrane but keep the actual proteins in your sample present. This means you can use a different antibody on the membrane to see if another protein is present. What you usually do is use GAPDH. This is your control antibody. And you use it to basically prove that everything has gone right. Because everything has GAPDH. GAPDH is strong so a concentration of 1:5000 is usually okay.  Stripping buffer is usually pre made and is located somewhere in the lab. Good places to find it are above the shaker or on the bench of the person who spends their entire life doing westerns.

35) Remove the PBS-T and add 5ml of stripping buffer.

36) Leave for 5 minutes at room temp on shaker. 

37) Wash again with PBS-T, 5 minutes each.  

38) Repeat steps 13 to 33.

39) Celebrate when you see your GAPDH bands.

40)Discard/ keep your membrane. Your choice. If keeping, leave it to dry, wrap it securely in clingfilm and put it in the -20 freezer. If discarding... well, just throw it in a bin. 

At best, it's been about three days since you started this. At worse, it's closer to a week. But congrats, you made it through your first western!  The first of many. I would like to say that you will start to enjoy it....  but you might find yourself resigned to it and stop hating western blots as much as you do at first. At least it feels absolutely amazing when it works. There is no feeling quite like it.