Western Blot

From Microbial Ecology and Evolution Lab Wiki
Jump to navigation Jump to search


To extract protein samples from E. coli, run them through an SDS-PAGE gel, and use antibodies to detect specific proteins in the size-separated protein samples. This technique is useful for confirming the presence of a particular protein inside or outside of E. coli, and for visualizing differences in protein expression across several samples.


Before Experiment

1. Autoclave however many Erlenmeyer flasks you need for the experiment

a. Number depends on how many E. coli variations you want to test (+1 for the negative control). For example, if you want to compare two strains of E. coli that are either uninduced (no target protein is theoretically expressed) or induced (target protein is theoretically expressed), you would need four flasks: uninduced strain 1, induced strain 1, uninduced strain 2, induced strain 2. And add one flask for the negative control: only LB broth.
b. To make sterile flasks, first grab unsterile flasks, cover the tops with aluminum foil, and autoclave them. Ideally, this should be done the day before Day 1 so that you have enough time to start and induce the cultures on Day 1, instead of waiting for the flasks to be autoclaved that same day

2. Set up liquid cultures of all E. coli strains you want to use. The general contents of each culture will be: 10 mL LB broth + x uL antibiotic (or none, depends on strain) + inoculate culture with freezer stock and incubate in the 37 degrees C shaking incubator at 125 rpm overnight. Antibiotic volume depends on antibiotic stock concentration and final volume in flask.

3. During the incubation step of the induced cultures in Day 1, you will need clamps to hold the flasks down onto the incubator floor since the rotation speed will be too fast for a sticky pad to hold the flasks down. Make sure you have enough, and the correct size, clamps to perform the incubation step. Rubber bands can be used in lieu of metal rings.

Day 1

Induction and Overnight Incubation of E. coli Culture

1. Grab however many sterile Erlenmeyer flasks you need for the experiment (including one for the negative control)

2. Label flasks with tape

3. Add x mL LB broth to all flasks

a. Volume used in MEE lab is typically 25-50 mL. Consult with Eric about the exact volume you would need.

4. Add <10 uL E. coli overnight culture and x uL of the corresponding antibiotics (if any) into the appropriate flasks

a. Antibiotic volume depends on antibiotic stock concentration and the final volume in the flask

5. Place flasks in the 37 degrees C shaking incubator at 125 rpm until the absorbance at 600 nm reaches 0.6-0.8.

6. Check on the cultures from time to time.

a. The most effective way is to get a 13mm test tube per culture you want to measure and another tube for a negative control. Add ~3 mL culture into the tubes, and add ~3 mL LB media into the negative control tube
b. Measure on the spectrophotometer at 600 nm and record the absorbances. The numbers will allow you to estimate how long you have to wait for the cultures to be done (e.g., abs = 0.2, and E. coli takes about 30-45 min to double, so you need to wait 60-90 min (two doublings) for the culture to be ready at 0.8).
c. Pipette the 3 mL of culture back into their respective flasks, making sure not to cross-contaminate
d. Put the flasks back in the 37C shaking incubator and check again later using the same test tubes

7. Screw in the appropriate clamps into the post-induction incubator before induction in order to make sure the induced cultures are immediately placed in the incubator—putting in clamps can take a while.

8. When the cultures have reached absorbance 0.6-0.8, add x uL inducer (e.g., IPTG) to the cultures that you want to induce

a. Volume of inducer depends on the inducer stock concentration, final volume in the flask, and the desired final concentration of the inducer. Multiple inducers may be added depending on the strain. 50 uM of IPTG (final) and 1.6 mM of 100% propionate (final) are ideal for inducing E. coli strains containing bacteriocin (IPTG) and transporter (propionate) plasmids.

9. Incubate all flasks for 12-18 hours overnight in the 34 degrees C shaking incubator at 200 rpm

a. 18 degrees C shaking incubator is typically used in other labs, but we found that 34 degrees C works just as well.

Day 2

Obtaining Supernatant and Lysate Samples from Bacterial Culture

1. Add the entire volume in each overnight flask to its own appropriately-sized falcon tube (don’t include negative control). Place tubes on ice. You don’t need to transfer culture in a hood; you can just transfer normally.

2. Use “fast temp” program on swinging-bucket centrifuge (in Bio Superlab) without adding any tubes to quickly cool the centrifuge down to 4 degrees C

3. Weigh the tubes to make sure they are within 0.1 g of each other (0.5 g at worst)

a. This step ensures the centrifuge is balanced

4. Centrifuge the tubes at 3900 rpm for 40 minutes at 4 degrees C

5. Transfer supernatant into another falcon tube and label appropriately. Keep on ice.

6. Resuspend the cell pellets in x uL lysis buffer by vortexing

a. To determine the volume of lysis buffer to use, use the following ratio: 15 mL lysis buffer per 800 mL culture
b. Lysis buffer recipe (5 mL total, adjust based on how much you need)
  • 150 uL 5 M NaCl (150 mM final) - chemical shelf
  • 50 uL 100% Triton X-100 (1% final) - chemical shelf
  • 1 mL 1 M Tris pH 8 (50 mM final) - chemical shelf
  • 3.8 mL MilliQ water
  • 50 uL beta-mercaptoethanol (1% final), MUST BE ADDED RIGHT BEFORE USING THE BUFFER, - flammable cabinet
  • 50 uL protease inhibitor cocktail, MUST BE ADDED RIGHT BEFORE USING THE BUFFER, - -20C freezer

7. Incubate the resuspended cells on ice for 10 minutes

8. Vortex cells for ~2 seconds to help break open the cell walls

9. Transfer 1.5 mL of lysate to a sterile 1.5 mL microcentrifuge tube

10. Centrifuge the tubes at 14000 x g for 30 minutes at 4 degrees C

a. Can use centrifuge in the coldroom or the one in the MEE Lab

11. Transfer all the supernatant (i.e., everything above the pellet) into sterile, labeled 1.5 mL tubes in preparation for SDS-PAGE analysis

Running an SDS-PAGE on Supernatant/Lysate Samples

1. Set the hot block to 95 degrees C to give it time to heat up. Fill enough wells with water to hold all protein sample tubes

2. Set up the BioRad gel apparatus. You may need to use a different type of gel depending on the protein of interest. The following gel apparatus section is specific to the BioRad gel.

a. Remove gel from package and REMOVE GREEN TAPE AT THE BOTTOM (failure to remove tape will result in gel not running properly)
b. Use a paper towel to dry the gel
c. Gently push the gel cassette out using your thumbs. Make sure the comb comes out evenly.
d. If running only one gel, grab a buffer dam (i.e., a “dummy” gel cassette)
e. Place gels and buffer dam in BioRad tetra system, making sure they fit snugly inside without falling out (have to make sure the cassettes are wedged inside the two half-circles at the bottom ledge). Also, MAKE SURE the tetra system has the electrode jacks that point out from the top—this is needed to run the current.
f. The gels must be on opposite sides of each other, with the lane numbers facing outwards. In the case of the buffer dam, which has no lane numbers, there should be text that indicates how to orient it (“gasket” = inside of the tetra system)
g. Close using the green clamps.
h. Place the tetra system in the gel box, with the black and red markers on the tetra system aligned with the black and red markers on the gel box
i. Prepare a 1X solution of Tris/Glycine/SDS running buffer, invert several times to mix thoroughly, and pour into the tetra system (NOT the whole gel box) until you fill it to the top without spilling over
i. How to make 1X solution: 200 mL of 10X running buffer in 1800 mL of MilliQ water. Adjust pH to 8.3
j. Lift the tetra system out of the gel box and check underneath for any buffer leakage
k. If there is no leaking, continue pouring running buffer until you reach just below the “blotting” line marked on the front of the gel box. Don’t go higher than this line since buffer will likely spill from the slits on the side.
l. If there is leaking, then the gel cassette and/or buffer dam are likely oriented incorrectly. Fix accordingly.

3. Prepare protein samples in loading buffer in 1.5 mL microcentrifuge tubes (volumes depend on the stock concentration of loading buffer and how much volume you can fit in an SDS-PAGE well). Pipette up and down to mix.

a. Also make sure to prepare a positive control and follow the manufacturer's protocol for how to prepare it.
b. NOTE: Preparing the protein samples in this way is technically not the "correct" method. Ideally, you want all protein+buffer samples to have equal concentrations of protein, since this will ensure that the SDS-PAGE gel runs evenly. To determine the amount of protein you need to add to the loading buffer in order to create equally-concentrated samples, you will need to run a Bradford assay. Ask the Cooke Lab (or another lab that performs Bradfords) for help with this.

4. Heat the protein samples at 95°C in the hot block for 5 min to unfold the protein

5. Quick spin samples for 5-10 seconds to get all the condensation down into the rest of the sample

6. Load x uL of protein ladder (this one is for 2-250 kDa proteins), the protein samples, the positive control, and, if necessary, a media-only negative control

a. Make sure to pipette up and down a few times before loading into the wells
b. Technique (with a P10 pipettor): Stick the tip against the front of the well, making sure you’re actually inside the well (there is a thin gap in which the tip must enter). Move the tip 3/4 down the well, then dispense without going to the 2nd stop in order to prevent air bubbles.
c. It is recommended to load 10 uL of sample at a time (i.e., 3 x 10 uL per 30 uL sample) so that the tip used is smaller and can fit in the wells better. Using a larger pipettor and pipetting all of the sample at once has been shown to cause spillage. Ideally, however, you will want to use a gel loading tip, which can dispense the entire sample at once without spillage.

7. Put on the electrodes/lid, making sure the cathode and anode on the lid match the ones on the tetra system

8. Run the gel for 1.5-2 hours at 70 V, constant voltage

a. Check on the power supply periodically to make sure the voltage is actually constant at 70 V; it can sometimes stray far away from the set voltage

9. Stop the current when the dye front reaches 4/5 of the gel

10. Remove the electrodes/lid, remove the tetra system, unclamp, and retrieve the gel cassette

11. Crank open the gel using a spatula at the indicated arrows along the sides of the gel cassette

a. Tip: start by inserting the spatula into the gap of the top-right arrow, lift up until that portion of the cassette breaks open a bit, then repeat with another gap further below, working your way toward the bottom-right arrow. Then, repeat starting with the top-left arrow, working your way toward the bottom-left arrow.

12. Slide the gel down into a small plastic container filled with tap water

13. Wash the gel in tap water for 5 minutes, with agitation, and repeat 2 more times (each time with fresh tap water)

Transferring the SDS-PAGE Gel to the Membrane

1. Open the Trans-Blot transfer pack

a. Video protocol
b. This transfer pack uses a 0.2 um PVDF membrane, which is better for low molecular weight proteins.
c. NOTE: this transfer pack requires a Trans-Blot Turbo Transfer System

2. Place the bottom stack (labeled “bottom”) on the transfer cassette—with the membrane facing up—and roll vertically and horizontally multiple times to remove bubbles

3. Carefully place the gel on top of the membrane. If any tears occur, use your finger to close the tear. Make sure that you align the gel with the membrane in such a way that all bands on the gel can be transferred to the membrane (i.e., don’t let a portion of the gel with bands hang outside of the membrane)

4. Roll again, making sure not to tear the gel

5. Place the top stack (labeled “top”) on top of the gel

6. Roll again

7. Close the lid of the transfer cassette and lock the lid by twisting towards the lock symbols

8. Insert the cassette into the transfer system

9. Click “Turbo”

10. Select “1 Mini TGX”

11. Use mixed MW program and select “Run A” or “Run B” (depending on which slot the cassette is in) to transfer for 3 minutes

12. Prepare 1X TBST solution in a 50 mL falcon tube

  • 5 mL 10X TBS
  • 45 mL MilliQ water
  • 50 uL Tween 20
  • Mix and shake

13. After the transfer is complete, remove the cassette, remove the lid, and throw out the gel and stacks into the regular trash (assuming the gel has no ethidium bromide or other toxic chemicals; otherwise, throw it out in the special container for hazardous materials)

14. Place the membrane/blot in a plastic container

15. Add enough 1X TBST to cover the blot, wrap in plastic wrap or cover with a lid, and store at -20 degrees C

16. Continue the blotting process tomorrow (assuming you do not have time to do it today)

Day 3

Primary Antibody Staining (Overnight)

This section's protocol is adapted from the official protocol, which is for a horseradish-peroxidase (HRP) substrate kit. However, you may need to use a different kit (and, consequently, a different staining protocol) depending on the desired protein sensitivity and the antibodies you're using.

1. Make Blocking Solution (5% Milk in TBST W/V or 2% BSA in TBST)

a. 5 g nonfat milk powder in 100 mL 1X TBST (5% nonfat milk Block)
i. This is ideal for the anti-FLAG M2 antibody, but another blocking solution may be better depending on the antibody you're using. Rest of protocol will be based on the use of this blocking solution
ii. Stir the blocking solution before use
iii. Keep refrigerated
b. 6.7 mL 30% BSA in 100 mL 1X TBST (2% BSA Block)
i. Another type of blocking solution that can be used instead of milk
ii. Keep refrigerated

2. Wash the blot with deionized H2O for 5 minutes with agitation to remove the TBST/transfer buffer leftover from yesterday’s transfer

3. Remove the water

4. Transfer the blot to a plastic container in which 10 mL of any liquid can entirely cover the blot. This is so that the 10 mL antibody solutions in which you will incubate the blot can completely cover the blot.

5. Block the membrane for 1 hour in 10 mL 5% nonfat milk block solution at room temperature with gentle agitation

a. In the meantime, prepare the primary antibody blocking solution in a falcon tube. Dilution below. Make sure to store antibody solution in 4C fridge if the blocking is not going to be done for another 30 min or more.

6. Remove the blocking solution by pouring it out over the sink

7. Incubate the membrane in a solution of primary antibody diluted in milk blocking solution, and leave it overnight at 4 degrees C with gentle agitation. Exact antibody dilution depends on the recommended values provided by the manufacturer of your antibody.

NOTE: If you are performing the Western blot to visualize the difference in protein expression between samples, then a loading control antibody is needed. This antibody typically targets housekeeping proteins that should have a constant concentration across all samples.

Day 4

Secondary Antibody Staining

This section's protocol is adapted from the official protocol, which is for a horseradish-peroxidase (HRP) substrate kit. However, you may need to use a different kit (and, consequently, a different staining protocol) depending on the desired protein sensitivity and the antibodies you're using.

1. Take the membrane from the 4 degrees C cold room and agitate for 5 minutes at room temperature

2. Remove the primary antibody solution

3. Wash the membrane three times in 10 mL 1X TBST (using new TBST for each wash), 5 minutes each. Use gentle agitation.

4. Remove the TBST

5. Incubate the membrane in a solution of secondary antibody diluted in 10 mL 5% nonfat milk blocking solution at room temperature for 1 hour. Use gentle agitation. Exact dilution depends on the antibody you're using.

6. Dump out the secondary antibody blocking solution in the sink.

7. Wash the membrane five times in 10 mL TBST (using new TBST for each wash), 5 minutes each with gentle agitation.

8. Remove the TBST

9. Transfer the membrane to a fresh container.

Imaging the Blot

1. Develop the membrane using the horseradish peroxidase (HRP) substrate kit (again, this is specific for certain antibodies/proteins) and the ProteinSimple FluorChem M imaging machine in Bio Superlab.

a. Pipette 5 mL of Stable Peroxide (white bottle) and 5 mL of Luminol/Enhancer (brown bottle) into a falcon tube. Invert to mix.
b. Pour the falcon tube contents over the blot. Incubate for 1-2 minutes.
c. Open the imager door, clean the black screen with ethanol, and place the black screen onto the imaging surface
i. Black screen is for chemiluminescent signals, you may need to use a different screen for different signals
d. Place the wet blot in a plastic cover (found in Superlab), push out bubbles, place onto the black screen, and close the imager door
e. Use the “Chemi with visible markers” setting, keep the rest of the settings on default
i. Setting depends on the tag used to visualize the protein. In this case, a FLAG tag requires the "Chemi with visible markers" setting
f. Click "expose"
i. Since most of the settings are on default, the blot will be exposed for an automated amount of time—this time can vary from blot to blot since it is based on how much time the imager “thinks” it needs to expose a particular blot. Time may have to be adjusted afterwards depending on the resulting image.
g. Label the image with your initials, the antibodies used, and the exposure time
h. Adjust the exposure time if the initial exposure did not show desired results
i. If the background is too strong, expose again, but for a shorter amount of time
ii. If no bands show up, resuspend blot in substrate solution for 1 min, and expose for a longer period of time
i. You can print out the image, but this typically reduces the quality. Ideally, you should get a flashdrive (in a drawer in the MEE lab, or your own) and insert it into the imaging machine in order to retrieve a high-quality image. Remove the flashdrive and insert it into your computer to transfer the image and add it to your lab notebook.

Final Note: You will definitely need to adjust this protocol based on your needs. Ask around for help with how to optimize conditions, volumes, materials, etc. for your experiment. Professor Amy Cooke (and her students), Professor Eric Miller, and Nicole Cunningham were very helpful in the creation of this protocol.