Lab 7: SDS-PAGE

1 Introduction

Protein electrophoresis is an extremely popular technique in molecular biology. Simply, proteins (typically from a cell or tissue lysate) in an SDS-containing buffer are added to the top of a polyacrylamide gel. The SDS, a powerful anionic surfactant, serves to surround the protein, overwhelming its inherent charge. The protein surrounded by the negatively-charged SDS has a net negative charge approximately proportional to its mass. When a potential difference is applied across the gel, the negatively charged proteins migrate through it. Smaller proteins migrate more quickly through the gel, and the proteins are separated by size into ‘bands’.

Sodium Dodecyl Sulfide – PolyAcrylamide Gel Electrophoresis (SDS-PAGE) is commonly followed by either total protein staining or transfer and Western blotting.


2 Objectives

- To separate proteins in a lysate by molecular weight.
- To prepare gels for total protein staining or Western blotting.


3 References

- SDS-PAGE Simulation
- The SDS-PAGE Hall of Shame
- Early Days of Gel Electrophoresis
- Image J


4 Reagents, Supplies, and Equipment

4.1 Reagents

1. 0.5 M Tris-HCl, pH 6.8

This buffer is used to making the stacking gel.

2. 1.5 M Tris-HCl , pH 8.8

This buffer is used to make the resolving gel.

3. 10% SDS (5 g of SDS in 50 mL dH₂O, pH 7.4)

4. 0.1% SDS (dilute 10% SDS with dH₂O)

5. Resolving gel materials (amounts for 2-3 6% acrylamide gels)

The resolving gel is the gel that is poured first; in it, proteins are resolved into discrete bands.

8.0 mL dH₂O
3.0 mL Acrylamide/Bis-Acrylamide
3.75 mL 1.5 M tris-HCl, pH 8.8
150 µL 10% (w/v) SDS
75 µL 10% (w/v) Ammonium persulfate (APS) in dH₂O, made fresh on day of use
10 µL TEMED

6. Stacking gel materials (amounts given for 2-3 4% acrylamide gels)

The stacking gel is poured on top of the resolving gel after it has finished gelling, with a ‘comb’. Protein samples are added to the individual wells formed by the stacking gel gelling around the comb.

3.0 mL dH₂O
666 µL Acrylamide/Bis-Acrylamide
1.25 mL 0.5 M tris-HCl, pH 6.8
50 µL 10% (w/v) SDS
5 µL 1% (w/v) Bromphenol blue
25 µL 10% (w/v) Ammonium persulfate (APS) in dH₂0, made fresh on day of use
5 µL TEMED

7. 5X gel-running buffer

This buffer is diluted to 1X with dH₂O to make gel running buffer. Gel running buffer is used to fill the electrophoresis cell (or bath), keeping the gel wet and allowing the electrophoresis unit to operate.

(5x)125 mM Tris base (30.3 g)
(5x)0.960 M glycine (144 g)
(5x)0.1% SDS (10 g)

Add the quantities in parenthesis above to dH₂O until total volume is 1800 mL, bring to pH 8.3, then add more dH₂O for final volume of 2000 mL. Store at room temperature and dilute to 1X with dH₂O before use.

8. 2X SDS-PAGE sample buffer

This is the buffer which protein samples are diluted with before being loaded into the gel for analysis.

9. Protein samples

10. Transfer buffer - Do NOT pH, make fresh on the day of use

The transfer buffer is used when transferring proteins from the gel that you poured to a nitrocellulose membrane (for Western blotting).

25 mM Tris base (3.0 g)
0.2 M glycine (15 g)
20% methanol (200 mL)

Add the quantities in parenthesis above and bring up to 1000 mL total volume with dH₂O.

11. Tris Buffered Saline with Tween-20 (TBST)

TBST is used to wash the nitrocellulose membrane.

20 mM Tris-HCl (3.14 g)
137 mM NaCl (8.0 g)
0.1% Tween-20 (1 mL)

Add the quantities in parenthesis above to 900 mL dH₂O, pH to 7.5, bring to final volume of 1000 mL with dH₂O. Store at 4°C.


4.2 Supplies

1. 1.5, 15, and 50 mL centrifuge tubes
2. 10, 100, and 1000 µL pipette tips
3. 5 and 10 mL pipette
4. Nitrocellulose membrane
5. Filter paper


4.3 Equipment

1. Mini-PROTEAN 3 Cell and Systems
2. Power supply


5 Protocol

5.1 Assembling the gel casting unit

1. Cover your lab bench with paper if you have not already done so.

2. Use distilled water to clean all electrophoresis equipment. Wipe with Kimwipe, and set the components out to dry.

3. Assemble casting stand as shown below.



4. Check assembly for leaks using distilled water. Fill the space between the glass plates; if the unit leaks, reassemble and try again. If not, pour the water out and dry between plates using a folded paper towel, Kimwipe, or filter paper.

5. Begin water boiling for later use.


5.2 Pouring the Resolving Gel

1. Mix all resolving gel components together except the 10% APS and TEMED in a 50 mL centrifuge tube. Seal tube and tip back and forth gently to mix.

2. Add 10% APS and TEMED, seal tube and tip back and forth gently to mix..

3. Transfer solution to each of the two plate sandwiches using a 5 or 10 mL pipette. Do not fill all the way! There should be ~1 cm of space between the top of the short plate and the resolving gel level (right in the middle of the green plastic bar behind the glass plates).

It is important that you add the correct amount of gel. Too little and your resolution will be poor; too much, and there won’t be room for a stacking gel on top. Be careful to avoid bubbles, which will inhibit polymerization and distort protein migration. Replace leftover gel in centrifuge tube.



4.Immediately after pouring the resolving gel, using a pipette or wash bottle slowly and very gently add just enough 0.1% SDS solution to cover the resolving gel without disturbing it. The SDS solution is there to keep the gel from drying out and to protect it from oxygen, which will inhibit the reaction.



5. Wait 15-30 minutes. Check to see that the leftover gel in the centrifuge tube has polymerized, and if it has, tilt the gel former slowly to confirm that the resolving gel under the SDS solution has polymerized completely.

Pour the SDS solution into a Kimwipe, and rinse the top of the gel very gently with dH₂O.



5.3 Pouring the Stacking Gel

1. Mix all stacking gel components together except the 10% APS and TEMED in a 15 mL centrifuge tube.

2. Add 10% APS and TEMED, seal tube and invert several times to mix. Add stacking gel solution to the top of the resolving gel using 5 mL pipette, returning extra solution to centrifuge tube.

The stacking gel solution should almost fill the remaining space – leave ~ 2-3 mm between the top of the stacking gel and the top of the short plate.

Immediately thereafter, insert the comb. Start from one side and 'brush' air bubbles off to one side.



3. Wait 5-15 minutes. Check to see that leftover gel in the centrifuge tube has polymerized.

5.4 Assembling the electrophoresis unit

1. After polymerization, remove your gel sandwiches gently from the gel casting apparatus and transfer them both into the electrophoresis unit as shown below.



When assembling the electrophoresis unit, the short plates must face inward!

2. Place the electrophoresis unit into the electrophoresis bath. Fill the inner chamber (the space between the two gels) with gel running buffer, and check for leaks.

3. Add approximately 250 mL of gel running buffer to the outer chamber (the clear plastic electrophoresis bath). 250 mL is about 4-5 cm of buffer, measured from the bottom. While the inner chamber must be filled, the outer chamber need not be completely filled.

4. Remove comb carefully and gently rinse each loading section of the gel gently with the gel running buffer using a 1000 μL pipette.

5. Set up equipment near power source.

5.5 Electrophoresis

1. Add 1 part 2X sample buffer to 1 part protein samples and standards.

2. Heat protein samples and standards (do not heat the molecular weight ladder unless the manufacturer suggests it) at 100 C for 5 minutes.

3. Put your samples on ice for ~60 seconds.

4. Centrifuge samples briefly to remove bubble and pellet any undissolved cell extract. Your protein will be in the supernatant; any pellet should be left undisturbed.

5. Using a 100 µL pipet tip, add protein samples to the wells in the gels as shown below. Be very careful! It’s easy to miss a well entirely if you’re not paying attention. Pipette very slowly.



The gel can be overloaded two ways - too much protein, or too much volume. Too much protein will separate badly; I suggest loading no more than ~40 µg of total protein. The maximum volume depends on the type of comb and the spacer plates used. For an 8-well comb using 0.75 mm spacer plates, I suggest using no more than 25 µL sample volume.

6. Run the gels at 100-150 V until the dye front has nearly reached the bottom of the gel (approximately 60 minutes).



5.6 Disassembly and Transfer

1. When finished, disassemble the electrophoresis unit.

2. Prepare a bath of transfer buffer (for a Western blot) and a bath of dH₂O (for total protein staining).

3. Use a wedge to gently separate one of the glass plates from the gels.

The gel might tend to stick at the top, where the stacking gel was. It doesn't matter which plate you remove, as long as you can remove one leaving the gel on the other. Be careful! It's easy to tear the gel at this point.



4. Carefully take a folded Kimwipe and place it atop the stacking gel (small gel on top with comb) without touching the resolving gel (larger gel on bottom without comb). The edge of the towel should line up with the separation between resolving and stacking gel. Press the paper towel firmly against the stacking gel, then pull the paper away.

The stacking gel should neatly come with the paper, and can be discarded. If it doesn't all come off at one, try again.

5. Submerge the gel in the bath as shown below. If necessary, use the wedge to loose the gel into the buffer bath.



5.6a Total protein staining

Typically total protein staining is done with Coomassie or silver stain. The protocol varies depending on the reagent that you use. This will be done next week.

5.6b Transfer for Western blotting

1. Cut a piece of nitrocellulose membrane and pieces of filter paper so that their size matches that of the gel. Try to avoid touching the nitrocellulose membrane, manipulating it only at the edges. After your gel has incubated in transfer buffer for ten minutes, add the membrane to the transfer buffer.

2. After your membrane has incubated in transfer buffer for five minutes, dip the filter paper and sponges into the transfer buffer. Using the mini trans-blot unit and the gel, make a sandwich of the following:

RED - sponge - paper - membrane - gel - paper - sponge - BLACK

Do not try to take the gel out of the bath without support. Slip either the membrane or the wedge underneath the gel, using it for support.



Don’t let the gel dry out, and pour a little extra transfer buffer over it if it's getting sticky and you're having trouble manipulating it.

3. Before sealing it, roll a pipette over the top paper layer of the sandwich to ensure that the membrane and the gel are pressed against each other without any air bubbles that will interfere with the transfer.

4. Insert this sandwich into the blotting apparatus and pour in enough transfer buffer to fill the transblot apparatus.

5. Run at ~100 V for ~ 60 minutes or ~25 V overnight in the refrigerator to allow transfer of proteins from the gel to the nitrocellulose.

6. Turn off mini trans-blot unit. Remove mini trans-blot sandwich, and carefully peel nitrocellulose membrane from gel. Wash briefly with TBST. Your membrane is ready to be blocked, probed with antibodies, and developed.