# SDS-PAGE

`PROTOCOL DUMP PLEASE FORMAT`

A. Solution Recipes

3.5X bis-Tris gel buffer

* 1.25 M bis-Tris in ddH2O
* pH 6.5-6.8 with HCl (around 10 mL 12 M HCl for 1 L buffer)

Acrylamide Solution

* 40% (w/v) acrylamide/bisacrylamide (37.5/1)
* ddH2O to volume

10% APS (make new APS each time you run a gel)

* 10% (w/v) ammonium persulfate
* ddH2O to volume

5X Low-MW Running Buffer (for separating 2-50 kDa proteins)

* 250 mM MES
* 250 mM Tris
* 5 mM EDTA
* 0.5% (w/v) SDS
* ddH2O to volume
* Add fresh before running gel:sodium bisulfite to 5 mM

Destaining Solution for Sypro Ruby-Stained Gels

* 10% (v/v) ethanol
* 7% (v/v) acetic acid
* ddH2O to volume

Resolving Gel Solution

* 1/3.5 volume of 3.5X bis-Tris buffer
* Acrylamide solution to 20% (w/v)
* ddH2O to volume
* To catalyze gel polymerization (DO NOT ADD until you’re ready to cast the gel!):
*
  * 25-30 μL 10% APS per gel
  * 6-7 μL TEMED per gel

Stacking Gel Solution

* 1X bis-Tris buffer
* Acrylamide solution to 4% (w/v)
* ddH2O to volume
* To catalyze gel polymerization (DO NOT ADD until you’re ready to cast the gel!):
*
  * 15 μL 10% APS per gel
  * 7 μL TEMED per gel

B. Casting the gel

![](https://lh5.googleusercontent.com/78lDVW1ytqn53d5m0LWpmNDETGE2NB_-pDqqszvwZsuqZOyWHYxxc5Nkm8EU7v3emX1NpJoYoMXqvSOmfesGwbaLDH5ILX5EBss9f_K5hs4n0vw-g6mpmLCSTIBRzKL7p72r84mC)![](https://lh4.googleusercontent.com/77yOlJy4CcvMx6tl3e6xGBQ0TYsu7yuOnpaoaZDA6whkQPXJuriSMjnyPiZqF8j2dGBDD3ljX45KxA0PRlP1Ax-aZbSZsZ0OCDH6eqZgwgLLzZQJEkzLtXhv7dvOMJKXxnFn2FNr)

```
           Gel casting stand      Gel casting frame
```

1. Place the gray gaskets into the holders on the bottom of the casting stand (Tip: you can coat the gaskets with petroleum jelly to make the seal tighter)
2. Place a short glass plate in front of a long glass plate (use the 0.75 mm plates). On a flat surface, insert the two plates into the gel casting frame so that the short plate is on the side with the clamps. Push the clamps to the side to clamp the glass in the frame.
3. Place the frame into one of the holders on the casting stand so that the clamps are facing outward.
4. Fill the space between the 2 glass plates with water to ensure that there are no leaks. Use a syringe with a gel loading tip and/or a piece of filter paper to remove the water.
5. Mix together the resolving gel solution (make 4 mL solution per gel) and use the syringe to cast the gel between the glass plates until the liquid level reaches the top of the frame's clamps (about 3-4 mL of solution).
6.
   1. Add the APS and TEMED quickly and swirl the solution a few times, then immediately cast the gel
   2. Rinse the syringe and tip with water immediately after casting the gel to prevent the gel from clogging the tip)
7. Quickly add a layer of bis-Tris buffer-saturated butanol on top of the resolving gel; fill to the top of the small glass plate. To make bis-Tris buffer-saturated butanol, take 2 parts butanol and one part bis-Tris buffer and shake together until emulsified. Let the mixture separate, then shake again and allow the layers to separate. Use the syringe to draw up the butanol layer.
8. After the resolving gel has set, use the syringe and filter paper to draw up and discard the butanol layer. Place the 0.75 mm comb halfway in (make sure the lettering on the comb is facing outward), leaving space to add the stacking gel solution.
9. Mix the solution for the stacking gel (make around 2 mL solution per gel) and use the syringe to add the solution to the top of the resolving gel. Stop when the liquid has reached the top of the short glass. Push the comb the rest of the way in, until the ridge on the comb rests on the top of the short glass.
10.
    1. Tip: add the APS and TEMED quickly and swirl the solution a few times, then immediately cast the gel
11. Once the stacking gel has set, carefully remove the comb

C. Loading and Running the Peptide Sample

1. Remove the glass plates and gel from the casting stand and frame, wiping off excess gel on the outside of the glass plates.
2. Assemble the electrophoresis chamber
3.
   1. Place the U-shaped gasket into the electrode assembly so that the notches on the tips of the gasket are facing outward
   2. Place the plates in the notches on the bottom of the electrode assembly with the small plate facing the U-shaped gasket (the notches in the gasket should rest on top of the small plate)
   3. Push the green side clamps in, holding the glass plates down so they aren’t lifted up when the clamp is pushed in
   4. Check the electrode assembly for leaks by pouring a layer of water into the center
   5. Remove the water and place the electrode assembly into the electrophoresis tank
4. Fill the center of the electrode assembly with1Xrunning buffer until the buffer covers the top of the gel but does not overflow into the electrophoresis tank
5.
   1. Tip: the power supply will read “Load Error” if the buffer has overflowed into the tank from the electrode assembly
6. Load 3 μL protein ladder to one of the gel lanes
7. Mix 5 μL sample with 5 μL 2X standard Laemmli Sample Buffer and add to gel lanes
8. Fill the electrophoresis tank with1Xrunning buffer until the liquid level reaches midway between the “2 gel” and “4 gel” marks on the tank
9. Run the gel at 100 V for 5 minutes, and then at 150 V until done
10.
    1. Tip: The bromophenol blue dye in the sample buffer runs around 3-5 kDa and can be used as an indicator of when to stop running the gel

D. Staining the gel

1. Remove the electrode assembly from the electrophoresis tank and discard the running buffer
2.
   1. Tip: the running buffer in the tank (but not from inside the electrode assembly) can be reused 3-4 times as long as the pH doesn’t change; do not mix used buffer with fresh buffer
3. Remove the glass plates and gel from the electrode assembly
4. Remove the gel from between the glass plates
5.
   1. Tip: Use the wedge tool provided with the casting kit to slowly pry off the small plate and lift one corner of the gel from the plate. Float the gel of the plate by inverting them under water and swirling gently until the gel separates from the glass
6. Rinse the gel with water to remove residual SDS. Rinse until bromophenol blue begins to fade slightly, but do not over-rinse or peptide could be lost

Staining with SYPRO Ruby:

1. Use a volume of Ruby stain that is 10 times the total volume of the gel(s)
2. Pour the stain into a polypropylene or PVC dish (do not use a glass dish)
3. Wrap the container in aluminum foil and put on a shaker or rocker at low speed
4.
   1. Tip: gels cannot be overstained with Ruby; stain overnight for the best results

Staining with Coomassie:

1. Pour enough stain into a staining dish to cover the gel(s)
2.  Place container on shaker or rocker at low speed for 30-60 minutes depending on how

    concentrated the peptides are; stop staining when bands are bright blue

E. Destaining the gel

Destaining SYPRO Ruby Gels

1. Put gels in around 100 mL destaining buffer
2. Wrap with aluminum foil and put container on shaker or rocker at low speed for 5 min
3. Change buffer (do not dump used buffer or Ruby stain down the drain!) and put container on shaker or rocker at low speed for 30 min
4. Gels can be placed directly on transilluminator for observation

Destaining Coomassie Gels:

1. Put gels in ddH2O and swirl several times; place on shaker or rocker at low speed for 5 minutes
2. Change water and swirl several more times; place on shaker or rocker at low speed for 30 minutes

F. Other Helpful Tips

* Liquid acrylamide is toxic, but once polymerized it’s harmless and can be thrown away in the trash
* Do not leave the gray gaskets from the casting stand to air dry after washing; pat them dry with a paper towel because they will degrade faster if left wet
