Knight:NuPAGE electrophoresis

For a much cheaper version of this protocol, see Sauer:bis-Tris SDS-PAGE, the very best.

Purpose

To run a denaturing protein gel. Note that a preferable version of this protocol is available at Sauer:bis-Tris SDS-PAGE, the very best (recommended by Kathleen). But since we seem to already have the Invitrogen kit contents in lab, this protocol describes use of the Invitrogen system.

Note that the gel and running buffer were chosen for a 13kD protein.

Materials

• NuPAGE Novex 10% Bis-Tris Gel 1.0 mm, 10 well
• XCell SureLock Mini-Cell electrophoresis apparatus
• 20X MES Running Buffer
• NuPAGE LDS Sample Buffer (4X)
• NuPAGE Reducing Agent (10X)
• NuPAGE Antioxidant
• SeeBlue Plus2 Pre-Stained Standard
• Gel drying solution
  • 20% ethanol
  • 10% glycerol
• Gel drying frames from Diversified Biotech
• Cellophane sheets from Diversified Biotech

Procedure

Running buffer preparation

Do this step first.

1. Prepare 1000mL 1X NuPAGE SDS running buffer
   • 50mL 20X MES Running Buffer
   • 950mL deionized water
2. Mix well.
3. Set aside 200mL buffer for use in Upper (Inner) buffer chamber).
   • Add 500μL NuPAGE Antioxidant immediately before running the gel.
   • Mix well.

Sample preparation

1. Wells can accommodate 25μL loading volume.
   • Likely they can accommodate more ... maybe 40μL.
2. Let sample buffer warm to room temperature.
3. Each sample should contain
   • 13 μL protein sample (max 0.5μg per band)
   • 5 μL NuPAGE LDS Sample Buffer
   • 2 μL NuPAGE Reducing Agent
4. Heat samples at 70°C for 10 mins.

Set up the gel apparatus during the 10 mins sample heating step.

Note that prestained molecular weight marker doesn't need any preparation.

Running the gel

1. Wear gloves.
2. Remove the NuPAGE gel from the pouch.
3. Rinse the gel cassette with deionized water.
4. Peel the tape from the bottom of the cassette.
5. Gently pull the comb from the cassette in one smooth motion.
6. Rinse the sample wells with 1X NuPAGE SDS running buffer.
   • Use a pipetman and pipet to squirt in running buffer.
7. Invert and shake to remove buffer.
8. Repeat rinse two more times.
9. Orient the two gels in the Mini-Cell such that the notched "well" side of the cassette faces inward towards the buffer core.
10. Seat the gels on the bottom of the Mini-Cell and lock into place with the gel tension wedge.
    • Use the plastic buffer dam if you are only running one gel.
11. Fill the upper buffer chamber with a small amount of upper buffer chamber running buffer (with antioxidant) to check tightness of seal.
    • If there is a leak, discard buffer, reseal chamber and try again.
12. Fill upper buffer chamber. Buffer level should exceed level of the wells. Requires about 200mL
13. Load samples.
14. Load protein molecular weight marker (20 μL per lane but 10μL also seems to work).
15. Fill lower buffer chamber at the gap near locking mechanism with 600mL NuPAGE SDS running buffer.
16. Run at 200V for 30 minutes.

Staining the gel

1. Shut off the power.
2. Disconnect electrodes.
3. Remove gels.
4. Insert a knife in between the two plates and pry the plates apart.
   • You should hear a cracking noise as you break the bond between the two plates.
5. Gently separate the two plates attempting to leave the gel on the bottom slotted plate.
6. Cut to separate gel from bottom lip.
7. Flip over and transfer gel to staining tray that has been prefilled with 100mL deionized water (see below).
   • Use lid of a 1000μL pipette tip box.

You have two options for staining at this point.

Fast, microwave protocol

1. Add 100mL deionized water in staining tray.
2. Microwave staining tray loosely covered for 1 min on high.
3. Shake staining tray for 1 min on orbital shaker at room temperature.
4. Discard water.
5. Repeat steps 1-4 two more times.
6. Add 20mL SimplyBlue SafeStain.
   • Need 100mL for staining tray.
7. Microwave staining tray loosely covered for 45 secs on high.
   • 1 min seemed a bit too long.
8. Shake staining tray for 5 mins on orbital shaker at room temperature.
9. Discard SimplyBlue SafeStain.
10. Add 100mL deionized water.
11. Add a kimwipe.
12. Shake staining tray for 10 mins on orbital shaker at room temperature.
13. Add 20mL 20% NaCl for at least 5 mins.
    • Only do this step if you aren't planning on drying the gel!
14. Gel can be stored in salt solution for several weeks.

This protocol seems to work pretty well.

Slow, room temperature protocol

1. Add 100mL deionized water in staining tray.
2. Shake staining tray for 5 min on orbital shaker at room temperature.
3. Discard water.
4. Repeat steps 1-3 two more times.
5. Add 20mL SimplyBlue SafeStain.
6. Shake staining tray for 1 hr on orbital shaker at room temperature.
7. Add 100mL deionized water.
8. Shake staining tray for 1 hr or more on orbital shaker at room temperature.

This protocol worked less well for me. However, I learned later that the typical trick people use is to include a kimwipe in the destain step to help soak up the stain. This will likely improve the signal to noise on the gel.

Drying the gel

1. Equilibrate gel in gel drying solution for at least 30 mins.
   • Reduces gel swelling and results in a more flexible dried gel.
2. Place two cellophane sheets in water for 1-2 mins.
   • Cellophane may appear cloudy but will clear upon drying.
3. Lay one sheet of cellophane on solid back plate, bevelled edge down. Avoid air bubbles.
4. Place gel on cellophane. Avoid air bubbles.
   • Air bubbles can cause cracking.
5. Pipet 1-2 mLs of gel drying solution on top of gel.
6. Layer a second wet sheet of cellophane on top of gel. Match edges with edges of back plate. Roll cellophane from bottom of gel towards the wells helps avoid air bubbles.
7. Place open frame over stack, bevelled edge up. Match edges of back plate. Frame should cover all edges of cellophane.
8. Attach plastic clips to all four sides.
9. Leave assembly to dry horizontally for at least 2 days.
10. Remove clips and pry apart assembly.
11. Peel dried gel/cellophane sandwich from back plate.
12. Trim off excess cellophane immediately to avoid curling.

Notes

• To date, the best staining tray I've found is the lid of a 1000μL pipette tip box. Then use a piece of mesh that just fits inside the lide to either keep the gel in place while changing solutions or to move the gel to and from the light box. This method requires smaller volumes of stain than the staining tray from Invitrogen.

Safety

• Use nitrile gloves when handling acrylamide.
• Dispose of acrylamide gels and trays as hazardous.

See here for detailed safety information.

References

1. NuPAGE Technical Guide