SDS-PAGE Protein Gels

Consolidated/Pictorial Protocol

SDS-PAGE sample prep and loading

Sample Prep - Marina Protocols

For whole cell protein SDS-PAGE

1. Grow cells to ~ OD 1.0

- For calculation purposes, you ideally want 1 ml of cells at an OD of 1.0

- If your samples are not at 1.0, increase or decrease volume to compensate (eg: If OD is only 0.747, use 1.34 ml)

2. Weigh eppindorf tube

- Need to subtract tube weight to determine wet cell weight (WCW) of sample -- don't forget this step!

3. Harvest appropriate sample volume and pellet cells

- RT benchtop centrifugation at 14K RPM; 2-3 minutes should be more than sufficient

4. Dry the pellet as completely as possible by decanting and then pipetting off any remaining liquid 5. Weigh dried pellet and eppindorf tube, subtract out tube weight and record WCW

At this point, pellet can be frozen and the following steps can be performed at a later date

6. Mix Laemmli sample buffer using the following recipe:

500 ul Bio-Rad 4x Laemmli buffer

450 ul dH2O

50 ul 2-mercaptoethanol

7. Add 100 ul of buffer to pellet and vortex to ensure cells and Laemmli buffer are well mixed

8. Incubate sample for 5 minutes at 95-100°C

9. Centrifuge for 5 minutes at room temp 15k rpm (benchtop centrifuge)

Samples are ready to be loaded into gel as described below

As of May 2013 5GB1 proteins are analyzed using Bio-Rad Mini-Protean TGX Precast Gels, 12% polyacrylamide (Cat# 456-1045)

For membrane-separated protein SDS-PAGE (and to isolate membranes for analysis)

1. Grow 50 ml cells to ~ OD 1.0

2. Weigh 50 ml tube and record

- Need to subtract tube weight to determine wet cell weight (WCW) of sample -- don't forget this step!

3. Transfer cells from vial to tube and pellet

4. Dry the pellet as completely as possible by decanting and then pipetting off any remaining liquid

5. Weigh dried pellet and tube, subtract out tube weight and record WCW

At this point, pellet can be frozen and the following steps can be performed at a later date

6. Resuspend cell pellet (should have 0.2-0.5g wet cell mass) in 3 ml of 50 mM Tris-HCl (pH 7.5) + 0.25M sucrose

7. French press at 1000 psi twice

8. Centrifuge at 5000 RPM for 10 minutes at 4°C (pellet will be mostly cell debris and mostly brown-pink)

9. Transfer supernatant to 50 ml screw-cap Nalgene tube and centrifuge at 14,000 RPM for 20 minutes (pellet will be white)

10. Weigh empty ultracentrifuge tubes and record

11. Transfer supernatant to ultracentrifuge tubes, balance carefully, and centrifuge at 100,000 x g (= 45K RPM) for 1 hour

12. Decant supernatant and dry remaining pellet well

13. Weigh pellet and tube, subtract tube weight and record collected membrane weight

• Collected membrane can be prepared for SDS-PAGE gel per directions above (Step 6)

Considerations for Loading the Gel

• Note that gels are both concentration- and volume-dependent; overloading of either will cause deformation of lanes and smearing
• 15 ul of 5GB1 prepared as above gives pretty good banding and a concentration ~4 μg/μL
• PageRuler Plus (Thermo Scientific) is a nice ruler
• Use BSA (usually comes at [10 μg/μL]) to make standards for calibration curve

A range of total BSA protein from 0.375 μg to 4.5 μg is a good place to start

Instructions for the quantification of protein bands can be found HERE

Running the Gel

• Bio-Rad Mini-Cell Setup
• Use diluted 10X Tris/Glycine/SDS Buffer

Re-using running buffer 2-3 times does not affect results

If only running one gel, don't forget to use buffer dam to replace second gel

• If using home-made gels, position so that the shorter plate faces inward!
• Apply pressure on gel holder and gels as you close the tabs to seal the center compartment

Mini-cell Gel holder

• Fill central compartment with running buffer

Be sure to pour enough to fill sample wells

• Pour more into the outer compartment to specified line
• Load gel

Make sure you will be able to determine the orientation of your gel after it is stained. Asymmetry is good!

• Run @ 60 V for ~15 min, then 200 V for ~ 20+ min

Note: ladder looks blurry while running through the stacking gel; don't be alarmed unless it still looks blurry in the resolving gel

Can skip the 60V step if you don't need a gorgeous gel

Amanda runs 20 min at 200V, then checks frequently to make sure the protein doesn't run off the gel

Storing Samples After Use

• Amanda was taught to flash-freeze (liquid N₂) samples and store them at -80^oC.
• Ladder is stored at -20°C. An aliquot of PageRuler Prestained that sat out at room temp for 1 week looked perfect in a gel, so don't worry about that ladder's stability.
• Nicole's experience is that more than 2 freeze-thaw cycles causes degradation in protein bands
• For best results do not save BSA samples for calibration curve; always make fresh standards!

Staining Gel for Visualization

1. Carefully remove gel from cast

2. Place in microwave-safe container not much larger than the gel

3. Pour Coomassie stain over gel, ensuring approximately 1/2" of coverage

• For quick staining, microwave container with gel and stain solution on high for 1 minute

- Make sure gel is completely covered by stain solution, or else gel can shrivel from the heat!

- Solution is smelly and methanol is toxic when inhaled; crack microwave door slightly and let steam dissipate before removing gel container

- Place container on orbital rocker and rock for 10 minutes

• For not-at-all quick staining, place container on orbital rocker and stain for 1 hour to overnight

4. Pour off stain solution into collection container (can be reused; refilter if necessary to remove gel chunks)

5. Rinse gel with dH2O to remove majority of remaining stain solution

6. Pour destain solution over gel, ensuring approximately 3/4" of coverage

• Add knotted chemwipes to greatly increase rate of detaining; be sure they don't stick to the gel and be sure to balance placement to ensure even destaining
• For quick destaining, microwave container with gel and destain solution on high for 1 minute

- Make sure gel is completely covered by destain solution, or else gel can shrivel from the heat!

- Solution is smelly and methanol is toxic when inhaled; crack microwave door slightly and let steam dissipate before removing gel

- Place container on orbital rocker and rock for 10 minutes

• For not-at-all quick destaining, place container on orbital rocker and destain overnight

7. Replace pigment-stained chemwipes and continue rocking, if necessary

8. Pour off destain solution into collection container (can be reused; store in a resealable container with chemwipe knots to pick up remaining pigment)

9. Visualize gel with equipment at hand, save image as .jpg or .tif; Instructions for the quantification of protein bands can be found HERE

Recipes

from/adapted from Joseph T.E. Roland [1]

Coomassie Stain - 1 L

0.1% Coomassie R250, 10% acetic acid, 40% methanol

Directions:

1. Add 100 ml of glacial acetic acid to 500 ml of ddH2O
2. Add 400 ml of methanol and mix
3. Add 1g of Coomassie R250 dye and mix
4. Filter to remove particulates (a coffee filter works great for this and is cheap)
5. Store at room temperature in a sealable container

• Stain can be reused numerous times
• Remove gel chunks by filtering through coffee filter again

Destain for Coomassie - 1 L

30% methanol, 10% acetic acid

Directions:

1. Add 100 ml of glacial acetic acid to 600 ml of ddH2O
2. Add 300 ml of methanol and mix
3. Store at room temperature in a sealable container