Western blotting

Natalya N. Pavlova   Craig Thompson Lab (MSKCC)

Western Blotting

Sample Preparation

Note: this method is for whole-cell lysis. It works for most cytosolic and nuclear proteins. Proteins that are tightly associated with chromatin require additional steps to break up chromatin - such as sonication or benzonase digestion.

1. Seed the cells in 6-well dishes, 300K-500K cells/well.
2. Carry out treatments as needed.
3. Once ready, place the dish on ice and thoroughly aspirate the culture medium.
4. Wash wells once with cold PBS.
5. Add cold lysis buffer to wells (100-150 μL per well)
   
   Lysis buffer (Note: prepare fresh; save a small aliquot as a blank for determining protein concentration):
   10x RIPA (cat. # 20-188, Millipore)                                 150 μL
   dH₂O                                                                              1350 μL
   100x protease inhibitor mix     (cat. #1860932)             15 μL
   100x phosphatase inhibitor mix (cat. #78428)               15 μL
    
6. Lyse cells on ice for 15 min; make sure the plate is level and rock the plate occasionally to redistribute the lysis buffer.
7. Scrape cells with plastic scrapers (one scraper per sample, wash between uses) into pre-chilled eppendorf tubes.
8. Spin down the samples at 4°C, 13,000g for 15 min.
9. Transfer the cleared lysates into a new set of eppendorfs, discard the pellet.
10. Mix a fraction of the lysate with 4x sample buffer (cat. #NP0007, Life Technologies) and 10x reducing agent (cat. #NP0009, Life Technologies), for instance: 65 μL lysate + 25 μL 4x sample buffer + 10 μL reducing agent.
11. Incubate at 95°C for 5 min. Watch out, as caps can occasionally pop.
12. Let the tubes cool to room temperature and spin down the condensation. At this point, samples can be stored indefinitely in -80°C, for several months in -20°C, or loaded on the gel the same day.
13. Use the remainder of the lysate to determine the protein concentration via BCA or Bradford assay according to the manufacturer’s instructions.
14. Using the information from #13, calculate the protein concentration of your samples.

Running the gel

1. Select your gel percentage and buffer type according to the size of the protein you are interested in resolving. E.g., 4-12% Bis-Tris gels are best for large and medium-sized proteins, 4-20% Bis-Tris are most suitable for smaller proteins and 3-8% Tris-Acetate gels are best for very large proteins. Make sure to use the correct running buffer for your gel (MOPS or MES buffer for Bis-Tris gels, Tris-Acetate buffer for Tris-Acetate gels).
2. Rinse the gel apparatus and gels with dH₂O. Remove the white stickers and combs from gels.
3. Assemble the apparatus and fill with 1x running buffer. One apparatus holds two gels, or you can use a dummy plastic cassette in place of the second gel.
4. Rinse the wells by vigorously pipetting 200 uL running buffer directly into wells.
5. Calculate how much volume of each sample you’ll need to load to have the same amount of protein loaded per well. Avoid loading less than 15 μg of protein per well. 20-30 μg is optimal.
6. Using gel loading tips, carefully load your samples into wells. Do not overfill! For the first lane, load the ladder (I use SeeBlue Plus 2, cat. #LC5925, Life Technologies).
7. Run gels at room temperature, 100V (constant voltage) until the blue dye front reaches the bottom of the gel.
8. Prepare the transfer buffer at least 2 hrs in advance and chill at 4°C (Note: the buffer is reusable up to 5 times):
   
   20x Transfer buffer  (cat. #NP0006, Life Technologies)       100 mL
   Methanol                                                                             200 mL
   dH2O                                                                                   1700 mL
    
9. Pre-soak the sponges in a dish filled with transfer buffer. Thoroughly squeeze out all the air bubbles.
10. Prepare cut pieces of Whatman paper pieces and nitrocellulose that match the size of your gel. Be careful not to touch the nitrocellulose with anything but a clean pair of forceps.
11. Disassemble the gel apparatus.
12. Open the gel cassette with a gel knife and carefully trim the thick part in the bottom and the wells.
13. With clean gloved hands, assemble the transfer sandwich. Keep everything submerged in a large dish filled with transfer buffer to avoid trapping air bubbles. Assembly order: Sponge-Whatman-Gel I-Nitrocellulose-Whatman-Sponge-Whatman-Gel II-Nitrocellulose-Whatman-Sponges (3-4). The whole sandwich should fit snugly into the transfer unit but not feel squeezed.
14. Insert and secure the transfer unit in a clean gel box and fill with cold transfer buffer.
15. Run at 40V for 2 hrs, room temperature.

Blotting

1. Disassemble the apparatus and stain the membranes with a few mL of Ponceau S (cat. #P7170, Sigma, reusable) for 1 min, rinse off the excess Ponceau with a couple of changes of 1x TBS-T and trim/cut your membranes as desired.
2. Rinse the membrane pieces in 1x TBS-T again until Ponceau comes off.
3. Block the membranes for 1 hr in 5% dry milk dissolved in 1x TBS-T.
4. Incubate the membranes overnight with antibodies of your choice dissolved in either 5% dry milk in TBS-T or 3% BSA in 1x TBS-T according to the manufacturer’s instructions. Antibody dilution for Western blotting is typically 1:1000 or 1:500, but may need to be optimized empirically.
5. Rinse the membranes briefly with two changes of 1x TBS-T, then wash in three more changes of 1x TBS-T, 10 min each.
6. Incubate the membranes for 1-2 hrs with a dilution of secondary HRP-linked antibody raised against the species of the primary antibody in 5% dry milk in 1x TBS-T. Secondary antibody dilution is typically 1:10000 or 1:5000.
7. Rinse the membranes briefly with two changes of 1x TBS-T, then wash in three more changes of 1x TBS-T, 10 min each.
8. Develop membranes in ECL solution (regular or super strength depending on the strength of the antibody signal) and image using either a digital or film-based method.
9. Membrane pieces can be consequently probed with a primary antibody against a different antigen of interest; however, steps may need to be taken to remove the signal from the first antibody first.

1. If protein of interest #2 is different in size from protein #1, the membrane can be simply rinsed in 1x TBS-T and put into antibody #2 dilution directly.
2. If protein of interest #2 is very close in size or is the same size as protein #1, and the primary antibodies targeting them are from different species, the signal from protein #1 can be quenched by incubating the membrane with 0.05% sodium azide in 1x TBS-T. This serves to inactivate the HRP enzyme activity from the secondary antibody attached to primary antibody #1.
3. If protein #2 is very close in size or is the same size as protein #1, and the antibodies are from the same species, the membrane can be stripped with the stripping buffer (cat. # 21059, Life Technologies) for 15 min and reblocked in 5% milk in 1x TBS-T for 30 min to remove the antibody bound to protein #1. The stripping method has limitations as it may not remove the signal completely or evenly and therefore must be used with caution.

Cautionary Note:

Verifying that the bands you see on your blot are veritably representing your protein of interest can be tricky and often requires extra work. Often, a mix of specific and non-specific bands of varying intensity is present; in addition, predicted molecular weight of protein of interest does not always match the empirical molecular weight in your particular cell line or tissue context. For phospho-specific antibodies, a positive control known to produce the change in phosphorylation is a good control (e.g. amino acid withdrawal to verify the change in pThr389 S6 kinase 1), while for antibodies directed against the “total” protein of interest, a genetic knockout or knockdown (e.g. via CRISPR, shRNA or a lysate from the knockout cell line) of your protein of interest is an ideal control.

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