Western Blot

Goal

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.

Protocol

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 gel apparatus (this is for the lysate samples)

a. Remove 8-16% 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 (this is the pH that Lou’s lab uses)

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 15 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

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