Western Blot

Sample Preparation: Cell Lysate and Conditioned Media

• Cells from 10 cm plates are are collected at 100% confluence, Day 0, in 1 mL of Western Lysis Buffer (WLB) and stored at -20 ̊C. Cells from 6 well plates should be lysed in 750 μL to 1 mL of WLB with protease tablets.

• Conditioned media is collected in 15 mL tubes, spun down at 5000 rpm for 5 minutes to form a pellet.

1) Cells can be treated with serum free (SF) conditions by washing with 1x PBS and adding 4-7 mL DMEM with 0.7% FBS or BSA.

2) 3T3-L1 cells at Day 0 should not be kept under these conditions for more than 6 hours, as cells begin to die at a rapid rate without serum.

3) Spin down the media collected from these cells since there will be cells suspended in it that must be cleared before proceeding.

4) Samples are boiled on a 95 °C heat block for 5 minutes (preadipocytes should be boiled for 15 minutes to fully disperse blob)

5) Pipet the sample to fully disperse the blob and remove an aliquot to use as a working sample.

6) 4x SDS loading buffer is added to the working sample to a concentration of 1x.

7) Samples are boiled again at 95 °C for 5 minutes to denature and allow for proper dispersal of the blob.

8) Working and stock samples should be stored at -20 °C.

Bradford Assay:

**Protein concentration can be obtained from the working sample immediately before addition if loading buffer (preferred) or subsequently from the stock sample. If concentration is determined from the working sample, remove 2 μl of the sample after the blob has been fully dispersed and add it to 1 ml of protein assay solution and proceed with the Bradford analysis. If the stock sample is being used, use the following procedure to determine protein concentration:

1. Preheat the stock sample at 60- 80 ̊C for 1-5 minutes.

2. Spin samples down for 1-2 minutes to cool them down and collect the sample that has condensed on the lid of the tube.

3. Pipet to fully suspend the blob

4. Remove 2 μl of the sample and add it to the 1 ml of protein assay solution and proceed with Bradford analysis.

Running the Gel:

1. Select performed gels based on the number of wells needed and set up the gel apparatus.

2. Once gel is firmly in place add running buffer to the center of the gel apparatus, between the gels, check for any leaks, then add Buffer to the outside of the gels about ½ or 2/3 the way up the apparatus.

3. Place the samples and run gel at 125 volts (constant) for ~90 minutes or until blue dye runs to the bottom of the gel.

Transferring the Membrane:

1. Fill a tray with 1x Transfer Buffer and place a transfer clip in the tray with the clear side down.

2. Prepare 3 pieces of Whatman paper cut to the size of the gel and one membrane cut a bit smaller for each gel.

3. Disassemble gel apparatus using a prying knife and trim off the stacking gel and the bottom part that has curled up.

4. Place a sponge down on the clear side of the clip, followed by two pieces of Whatman paper, then the gel.

5. Prewet the membrane in the transfer buffer for a few seconds then place the membrane on top of the gel.

6. Add another piece of Whatman paper, then the other sponge and close transfer clip.

7. Place transfer clip in transfer apparatus so that the black side of the clip is facing the red side of the apparatus. Proteins are negatively charged because of the SDS and will migrate to the positive (red) charge.

8. Pour the 1x transfer buffer in the tray into the transfer apparatus and add any additional buffer needed to fill the apparatus to the top. Place the lid on and run at 35 volts overnight. Alternatively the transfer can run at a higher voltage for a few hours, but an ice block must be placed in the transfer apparatus to prevent overheating.

Blocking the Membrane:

1. Prepare 5% milk and 1x TTBS. 10x TTBS is a stock solution in the cold room. Dilute to 1x with Milli-Q H2O and use for the remaining steps (usually about 500 ml is needed). 10 ml of 5% milk is needed for each blot when blocking.

2. Cut a bag that is larger than the blot as adjustments may be needed once the bag is sealed.

3. Disassemble the transfer apparatus and remove the membrane. Cut the membrane corner (bottom corner opposite side of the marker) to orient the membrane later on. Discard the gel and the Whatman paper, and rinse the rest of the transfer apparatus and let dry.

4. Immediately place the membrane in bag and seal the three sides. Add 10 ml 5% milk for each membrane/bag and steal the remaining side of the bag making sure not to seal in any bubbles.

5. Place the bag on top of the rocker with a large bag of water on top of it and block in 5% milk for 1 hour.

6. Remove the membrane from the bag and place in a tip box with enough 1x TTBS to cover and wash 3 times on rocker for 10 minutes each time.

Primary Antibody Incubation:

1. Prepare primary antibody by diluting in either 5% milk or 1% γ-globulins in 1x TTBS. For the anti-myc-HRP antibody, dilute 1:1000 in 1% γ-globulins in 1x TTBS (0.1 g in 10 ml).

2. Cut a bag that is larger than the membrane.

3. Remove the membrane from the tip box and trim any unneeded parts of the membrane off then place the membrane in the bag and seal three edges.

4. Add 5 ml of the primary antibody solution to the bag and seal without any bubbles. Let rock with a large bag of water in top of it for 2 hours at RT. (Can go overnight at 4 ̊C)

5. Remove membrane from bag and wash 3 times for 10 minutes each in 1x TTBS.

Secondary Antibody Incubation:

1. During final wash, prepare second antibody by diluting in BSA or non-fat dry milk

2. Cut a bag that is larger than the membrane

3. Remove membrane from tip box, place in bag and seal three edges.

4. Add 5 ml of the secondary antibody solution per membrane/bag and seal the final side of the bag. Incubate on the rocker with a large bag of water on top for about an hour at RT.

5. After incubation, wash membrane in a tip box with 1x TTBS 3-4 times for 10 minutes each.


1. During the last wash, mix ECL reagents 1:1 (for 1 membrane, mix 3 mL of each reagent and add 1/10 volume enhancer reagents (300 μL of each) in the bottom of the tip box.

2. Remove membrane from bag and expose to ECL mixture in tip box for about 2 minutes.

3. Place membrane in plastic wrap and tape plastic wrap to the inside of a film cassette.

4. Expose film to membrane in dark room and develop film.