Total Protein Extraction and 2-D Gel Electrophoresis Methods for Burkholderia Species

1. Culture Growth (Days 1+2)

1. Grow a starter culture: 3 ml of Luria Bertani (LB) broth in a snap top 15 ml tube, at 37 °C rotator overnight. Dilute overnight growth to an OD₆₀₀ of 0.6. Add 1 ml of this dilution to 100 ml of LB broth in a 250 ml Erlenmeyer flask. Incubate at 37 °C with shaking at 250 rpm for 16 hr into stationary phase (SP), to better approximate, in batch culture, infection conditions and to assure that bacterial virulence factors under the control of stationary phase signal factors such as rpoS and quorum sensing, are expressed. Alternatively, bacteria grown at early SP or mid- or late-logarithmic phase can be used to gain information on the bacterial adaptation phase.
2. After 16 hr measure the OD₆₀₀ and ensure the culture has reached SP by comparison to a previously constructed growth curve under identical conditions.

2. Protein Extraction (Day 3)

1. Chill all media and solutions beforehand in fridge overnight. All steps should be carried out with tubes in an ice bucket. Prechill a microcentrifuge and a Beckman Coulter High Performance centrifuge (or equivalent) to 4 °C and the rotor inside (rotor=JA-20, centrifuge=J2-HS).
2. Prepare fresh 0.1 M phenylmethanesulfonyl fluoride (PMSF, 174.19 g/mol) solution in acetone by dissolving 0.0174 g of PMSF ('CAUTION' PMSF is toxic and corrosive, use face shields and gloves) in 1 ml of acetone ('CAUTION' acetone is toxic and flammable, use face shields and gloves).
3. Make up a PBS/EDTA/PMSF solution: 0.1 ml of 0.1 M PMSF, 0.1 ml of 0.5 M EDTA, and 9.8 ml of PBS. Check the OD of the 16 hr culture is approximately where it should be in SP.
4. Take 35 ml of the bacterial culture and transfer to an Oakridge centrifuge tube and centrifuge the culture at 4,500 × g for 20 min at 4 °C.
5. Remove supernatant to a waste container and add 35 ml of cooled PBS and resuspend the pellet. Centrifuge again at 4,500 × g for 20 min at 4 °C.
6. Remove supernatant and resuspend in 1 ml of cold PBS. Transfer to sterile 2 ml Eppendorf and centrifuge 1 min on high in the microcentrifuge at 14,000 × g at 4 °C. Remove and discard supernatant.
7. Add 1 ml of PBS/EDTA/PMSF solution, resuspend the pellet and transfer to a 2 ml screw cap tube (with o-ring) containing ~0.5 ml glass beads (presterilized). Add the re-suspended pellet to the tube containing the glass beads and bead bash them at 4 °C for 1 min in cold-room. Do this 3x, putting the sample on ice after each bash.
8. Centrifuge at 14,000 x g for 1 min in the microcentrifuge. Remove the supernatant and put it in a sterile 5 ml polystyrene tube.
9. To increase the yield add 1 ml of PBS/EDTA/PMSF solution to same tube contain glass beads again. Bead bash the tube at 4 °C for 1 min. Do this 2x putting the sample on ice after bash. Centrifuge at 14,000 x g for one minute on the centrifuge and remove the supernatant and add to supernatant from step 2.8. Use a 1 ml syringe and a 0.20 μm filter the supernatant from the polystyrene tube to a new sterile 2 ml Eppendorf tube.
10. At this point the samples should be assayed for protein quantity, using the Pierce MicroBCA protein extraction kit and aliquots of 200 μg should be frozen at -80 °C until required.

3. Sample Preparation (Day 4)

1. Prepare 25 ml rehydration stock solution (8 M urea, 2% CHAPS, 2% IPG Buffer, 0.002% bromophenol blue) and store in 2.5 ml aliquots at -20 °C. Add 7 mg dithiothreitol (DTT, 154.2 g/mol) just prior to use to a 2.5 ml aliquot of rehydration stock solution.
2. Process thawed samples from step 2.10 using a 2-D Clean up kit with final resuspension in 450 μl of rehydration stock solution prepared in step 3.1.

4. First-dimension Isoelectric Focusing (IEF) (Day 4)

All steps in this section are using the Ettan IGPhor IEF System.

1. Set up the IPG first dimension strips: clean strip holders with strip cleaning solution and then wash with Milli-Q water, leave upside down to dry. Assign each sample to a gel strip holder number.
2. Load the sample from rehydration step in the second wider area from the (-) end. Use clean forceps to pull gel strips out of packaging. Take out protective layer from gel strips. Line strips rough side down in a slow, sliding motion to get the strips wet along the way from (-) end to (+) end.
3. Put damp blotting paper with sterilized water on top of electrode and under the gel strips to prevent burnout. Rinse tweezers in-between rounds with purified water and ethanol. Remove all bubbles, overlay thinly with mineral oil to prevent dry outs. Line the gel strip holders on the machine.
4. Run 12 hr at 20 °C for rehydration, 1 hr at 200 V, 1 hr at 500 V, 1 hr at 1,000 V, 0.5 hr at 4,000 V, 12 hr at 8,000 V, 24 hr hold at 300 V and can be taken out at this time.
5. After IEF is finished, it is recommended to perform the second-dimension SDS-polyacrylamide gel electrophoresis, unless the IPG strip are being frozen at -80 °C in Saran Wrap coated with mineral oil for future analysis. Alternatively, in order to avoid point streaking due to DTT access it is possible to include a second equilibration step of the IPG strips prior to the second dimension with a buffer that contains iodoacetamide.

5. Second-dimension SDS-polyacrylamide Gel Electrophoresis (Day 5)

All steps in this section are using the Ettan DALTsix electrophoresis apparatus.

1. To assemble the gel caster apparatus, thoroughly clean all components and ensure that gel caster is level and the grey rubber binding is lubricated with silicon gel. Put the black rubber triangular stopper on the bottom. Then, alternately place the separator and the glass plates with the shorter plate of glass facing front. Make sure the separators are on the back and the front. Fill the remaining space with the fillers (about 5 sheets) so the top is flat. Put the front panel on with the red tip on the outside. Use clamps to tighten each side and tighten the bottom screws.
2. Preparing the gel: Add 297.3 ml of Duracryl, ('CAUTION' Duracryl is toxic use face shields and gloves), 147 ml of Tris Buffer 1.5 M pH 8.8, and 143.3 ml of Milli-Q water into a flask with vacuum tip containing a stir bar. Put lid on top and mix solution at moderate speed with the vacuum on for 20 min. The vacuum is used to de-gas the solution. Make a fresh 10% of ammonium persulfate solution (APS, 218.8 g/mol): 0.2 g of APS to 2 ml of Milli-Q water.
3. Adding the gel mixture to the gel caster: When vacuum is done add 6 ml of 10% sodium dodecyl sulfate (SDS, 228.38 g/mol) to the side of the flask to avoid putting more bubbles in the solution. Add APS, stir, add 0.25 ml N,N,N',N'-tetramethylethylenediamine (TEMED, 116.20 g/mol) ('CAUTION' TEMED is flammable use face shields and gloves) and stir.
4. Pour gel mixture into gel caster through a funnel inserted at the back of the apparatus. Pour up to an inch below the short plate. Add 1.5 ml of water-saturated butanol to the top of the gel and wrap top with saran wrap to avoid dehydration. Wait for 1 hr to ensure full polymerization.
5. Checking the gel: When the gel is done (check the flask), disassemble the gel caster by a sink. Check for gel integrity while rinsing the glass plates with warm water; do not allow water to enter the plate. Pour out the butanol and rinse the top of gel twice with water to get rid of butanol. Fill the top with water again and let sit if the gels are not used right away.
6. Preparing the strips: Prepare equilibration buffer solution from the freezer (6 M Urea, 75 mM Tris-HCl pH 8.8, 29.9% SDS, 0.002% bromophenol blue). This can be preprepared and stored in 10 ml aliquots at -20 °C. Dissolve 0.1 g of DTT to a 10 ml of equilibration buffer solution. One tube can be used for two IPG strips. Place the gels of the strip face down onto the buffer and let sit for 30 min. Remember which end is which.
7. Preparing the Electrophoresis unit (2-D separation apparatus): Add 4.5 L of 1x electrophoresis buffer (25 mM Tris Base [121.1 g/mol], 192 mM Glycine [288.38 g/mol], and 0.1% w/v SDS [288.38 g/mol]) to the running tank. This buffer can be used for up to 3x. Turn the electrophoresis unit on. Check water level in water-cooling machine and fill it up with water if it is low. Turn the machine on, press mode to check the designated temperature, which should be 10 °C.
8. Make 1 L of 2x running buffer and 1 L of 1x of running buffer and store them at 4 °C. Make agarose-sealing solution: weight 0.25 g of agarose and dissolve in 50 ml of 1x running buffer. Pulse for 15 sec each.
9. Setting up the Electrophoresis unit: Use tweezers to take out each strip and remove the excess buffer on both sides with clean paper towel. DO NOT touch the strips. Pour off the water on top of the gel and line the gel top with 1x running buffer. Use clean tweezers and place the strips on top of the long plate, with the gel side facing towards you. Add 1x buffer to the strips to lubricate it. Slide strips further down by using plastic strips. Remove all bubbles in between the strip and the gel.
10. Add agarose-sealing solution to the top of the strip to seal it. Repeat for the rest of the strips. Place the glass plates containing the gels into gel cradle and lower into the gel tank. Ensure the 1x buffer level in the outer chamber is on the designated bottom line before putting the upper chamber on.
11. Place the frame for upper buffer chamber on top of the glass plates and ensure it is all the way down by pressing the bottom down. Use a funnel and add the 2x running buffer into the upper chamber up to the middle line. Use a funnel and add 1x running buffer to the outer chamber until it reaches the top line. Place lid on top and turn on the Electrophoresis unit and the power pack.
12. Run overnight at 52 V, 96 mA, 5 W. Check if the current is going by looking for bubbles on the silver wire near the top. Run the gel until the leading line is 1 cm from the bottom.
13. Proteins can also be analyzed using equipment and reagents from other companies such BIORAD or Hoefer, according to manufacturer instructions.

6. Silver Staining of the Gels for Gel Visualization (Day 5-6)

1. Prepare a fix solution 1 (800 ml ethanol, 200 ml acetic acid, and 1,000 ml of Milli-Q water in fume hood into a plastic bucket). Solution is good for 6 gels. Turn all the machines off and take out the whole middle section of the Electrophoresis unit and pull it in sink. Also put the white stand in the sink. Pull out the top tray containing the 2x running buffer.
2. Disassemble the apparatus and remove the gels from the glass plates into fix solution 1. Gels can be identified by cutting off corners when they are removed from the glass plates, the number of corners cut corresponding to the gel order in the caster.
3. Place the container with the fix solution and the gels on a rocker for at least 1 hr to overnight at 4 °C.
4. Transfer the gels to fix Solution 2 (20 ml of 50% glutaraldehyde, 600 ml of ethanol, 5 g of potassium tetrathionate (302.46 g/mol), 136 g of sodium acetate (82.03 g/mol), and Milli-Q water to 2,000 ml) and place on a rocker for 1 hr.
5. Wash the gels 4x for 15 min each in Milli-Q water.
6. Stain the gels in Silver Nitrate Solution (4 g of silver nitrate (169.87 g/mol), 500 μl of formaldehyde, and Milli-Q water to 2,000 ml) for 30 min or up to 48 hr.
7. Wash gel for 1 min in Milli-Q water.
8. Transfer to developer Solution (60 g of sodium carbonate (105.99 g/mol), 300 μl of formaldehyde, 15 mg of sodium thiosulfate (158.11 g/mol), and Milli-Q water to 2,000 ml) for 5-30 min until the gel is stained.
9. Put the gels in Stop Solution (100 g of Tris-Base (121.14 g/mol), 40 ml of acetic acid, and Milli-Q water to 2,000 ml) for 10 min.
10. For storage, transfer the gels to Glycerol Solution (40 ml of glycerol or 400 ml if long term storage is desired, and 1,600 ml of Milli-Q water) for 10 min and then dry them using a gel dryer.
11. Gels can be visualized after staining using imaging systems such as a scanner/densitometer, photographed using light transilluminator or Gel imager. Image Analysis software such as PDQuest 2-D analysis from BIORAD can then be used to obtain quantitative and qualitative information from proteins in a sample.

7. Gel Silver Staining for Mass Spectrometry Analysis

1. Fix the gels in 50% methanol, and 5% acetic acid for 1 hr.
2. Wash in 50% methanol for at least 10 min to overnight.
3. Wash in distilled water 3x for 10 min.
4. Sensitize with 0.02% sodium thiosulfate (158.11 g/mol) 2x for 15 min.
5. Wash 3x in CHILLED distilled water 3x for 10 min.
6. Submerge gel in CHILLED 1% silver nitrate solution (169.87 g/mol) for at least 20 min to 1 hr at 4 °C
7. Wash 2x for 1 min with distilled water. Change tray/bucket.
8. Develop in 0.04% formaldehyde in 2% sodium carbonate anhydrous (105.99 g/mol) and discard when it turns yellow quite quick; have three batches and be ready to change every 5 min.
9. Wash in 5% acetic acid then store in 1% acetic acid.
10. Rinse 1.5 ml Eppendorf tubes (to hold get spot cut-outs) with USP grade ethanol and air-dry prior to use.
11. Clean forceps, razor blades and surfaces with 100% methanol.
12. Spray gloves and surfaces with ethanol.
13. Excise spots in a flow hood or clean air area. Spots can be also excised using an automated spot cutter.
14. Put them in Eppendorf tubes and ship on ice or keep in fridge.