DAY 1 - Preparation of Samples and Gels
1. Collect a cell pellet (minimum 50,000 cells, this much will give you enough whole cell lysate (WCL) for 1 western if lysed with 25uL of Modified Lysis

     Buffer, 4 uL will be used for quantification leaving you with approx. 20uL of lysate for loading, if you load 5-10ug maybe you can get 2 westerns out

     of this many cells)
2. Depending on pellet size lyse with anything from 50uL to 500uL of lysis buffer.  Pippette up and down several time to properly lyse the cells
      a. A good rule of thumb is if the pellet is from a 6well plate use 50-100uL lysis buffer. If from a 10cm dish use 100-200uL lysis buffer.
3. Place the whole cell lysate into 1.5ml tubes if they are no in them already. Put the 1.5mL tubes containing the WCL in a tube rack. Put the tube rack  

     on a Newtater at 4degrees for 20minutes
4. Remove testube rack from 4degree and centrifuge 10min @10,000 RPM.

    a. Make sure you use a cooled centrifuge, the temp should be set to 4deg
5. Obtain new 1.5mL tubes, label them accordingly. Transfer WCL into them.
    a. After centrifugation you will notice a large pellet at the bottom, this is the insoluble fraction. The larger your original pellet was the larger the

        insoluble fraction will be, and vice versa.
6. Do a protein quantification assay for your samples.
    a. We use the Pierce BCA Assay Cat# 23225
7. Here you can do one of two things:
    a. Take all your lysate and standardize it to 1ug/ul or [whatever] you like using a combination of 3X and 1X Loading dye. The benefit of doing this is

        that it allows you to quickly run westerns and never re-quantify your samples. The downside of doing this is that once you standardize your

        samples you will disrupt ALL protein-protein interactions preventing you from looking at protein interaction within your lysate.
    b. Make fresh samples every time, this will be described in more detail.
8. After quantification calculate the amount of lysate you need. I will use an example here. Lets say your lysate contains a protein concentration of

    5ug/ul and you want to load 20ug. 20ug/5ug/uL = 4uL of lysate. To this you will then add 2uL of 3X Loading Dye (Recipe is provided in protocols). You

    do this because your lysate will dilute the 3X into 1X. Then you will top up to with 1X Loading Dye up your desired volume. Recommended volumes

    are 5-20uL if loading into a 15well gel or 10-50uL if loading into a 10well gel. Keep your samples on ice as you prepare them.
9. Boil your samples on a heat block (80deg-100deg) for 5 minutes. This will ensure everything is denatured and that the SDS in your loading Dye will

    nicely coat your proteins with negative charge.
     a. When boiling you can use lid-locks or you can risk it and put a heavy object on top of your 1.5mL tubes. If they pop, you will have to remake them.

          Lid-locks are recommended for precious samples….
10. Spin down your boiled samples. Use the quick spin option on the centrifuge until you reach approx. 9kRPM
11. At this point you can store your prepared samples for 1-3 days at -20degrees.
12. Your WHOLE CELL LYSATE should be stored at -80degrees.
13. Make desired % SDS gel and use desired comb for number of wells.

Store your gels by wetting paper towel with either deionized water or 1X Running buffer and wrapping them with this paper towel. Then stack a bunch of gels (wrapped in paper towel) together and wrap them in saran wrap. Place this at 4 degrees.

DAY 2 - Running Westerns, Transfer, Blocking, Primary Antibody
1. Start early in the morning as this is the labor intensive day.
2. Take our your gels, take our your samples, make 1X running buffer (recipe provided in protocols), get a protein ladder of your choice.
3. Set up the westernblot apparatus, and run the gels.
4. Initially you should run at approx. 80Volts to ensure that all samples stack nicely in the stacking gel. After about 20-30minutes you can increase

     voltage to 120-130Volts. Going higher is risky, as you will cook your gel.
5. Running your gel can take anywhere from 45min-2hours depending on %acrylamide use to make your gel.
     a. While your gel is running you should activate the PVDF membrane by Newtating it in methanol for approx. 2 min. After the 2min are up, discard

          methanol, do a rinse in ddH20, and then fill the container up with 1X Transfer buffer. Leave PVDF in transfer buffer until you are ready to use

          them. For larger proteins you can use 0.45um pore sized PVDF for smaller proteins use a smaller pore size PVDF membrane.
               i. Just for pure interest: Methanol opens pores in pores in your PVDF allowing aqueous substances to pass through the PVDF and interact with 

                 whatever is bound to it, in this case it will be your proteins. As your antibodies will be diluted in polar solutions (ie: water) you need your

                 PVDF activated.
6. After running, take out your gel, pry the two glass plates apart GENTLY and place the gel onto an ACTIVATED PVDF membrane. Newtate the gel and

    PVDF membrane for 5 min to qualize the gel and saturate it with 1X transfer buffer.
7. Use a Semi-Dry transfer apparatus from BIO-RAD.
     a. http://www.bio-rad.com/prd/en/BN/LSR/PDP/LGOQBW15/Trans-Blot-Turbo-Transfer-System.
     b. This machine is so worth getting it.
8. Take out a cassette, open it, pour 1X transfer buffer inside, be generous. Place a 1 thick wattmann filter paper on the bottom, if using a thin wattman

     paper use 3. Place PVDF and gel on top. Place another thick wattman paper on top of this. The sandwich should be, from top to bottom,

9. Set transfer machine to these settings. This works really well for a 10% gel
     a. Voltage – 10V
     b. Current – 0.2Amps
     c. Time – 65 minutes
10. After transfer is complete, block for 1+ hours in 5% skim milk/TBST or 5% BSA/1XTBST. (Recipe for making TBST are in the protocols section)
     a. You can block for 1h+ at RT or 1-5days at 4degrees, just make sure you cover the container with saran wrap to prevent your blocking solution from evaporating.
11. After blocking do a rinse with TBST
12. Add primary antibody at a desired concentration in 5%BSA/TBST or 5%skim milk/TBST.
    a. How to save antibodies. A good rule of thumb is that if an antibody is against a covalent protein modification such as a phosphorylation,  

        methylation, and so on it may not be reusable. Nevertheless you should still try reusing it. Otherwise, the antibody can be reused.
    b. To save antibodies you will have to use 5% BSA (with either PBST or TBST). After probing PVDF, do not discard the antibody. Place it in a 10mL tube

         add Sodium Azide to a final concentration of 0.05% and store at either -20 or 4degrees if you use the antibody often. Some antibodies lasted me

         for several months. Also if your antibody is VERY DIRTY the first few times, reusing it may clean it up.
c. Primary antibody can be incubated overnight at 4degrees or RT for 4-5hours.

DAY 3 - Washing, Secondary Antibody, Exposing Film
1. Discard the primary antibody or save it if you want to.
2. Do 4x15min washes with TBST
a. If this is your first time using an antibody and you want a VERY clean blot wash 4x25min or 5x25min. This will make sure you get a CLEAAAN blot.
3. Add secondary antibody, usually a concentration of 1:5000 works. Incubate at room temp for 1h
4. Once again, wash 4x15 times.
5. After the last wash discard the TBST and add either TBS or PBS (doest matter). Then proceed to expose the film.



Click here to download the protocol

By: Yev Chornenkyy