SDS-PAGE Protocol

[moleculardude]

SDS PAGE (Polyacrylamide Gel Electrophoresis)

Materials:

* Molecular Weight Marker
* 4-20% acrylamide gradient gels
* Tris-glycine-SDS buffer
* Practice gel loading solution
* Marker protein
* Sample proteins
* Sealant
* 50 ml Coomassie Blue Staining Solution
* De-staining solution (7.5% acetic acid)


Equipment:

* Pipette and tips
* Gel machine
* Staining box
* Hot water bath for boiling samples
* Microfuge
* Saran wrap


Background:
Gel electrophoresis is a technique in which an electric current is used to move charged molecules through a gel-like matrix. The direction, distance, and speed of migration are dependent on the size, shape and electrical charge of the molecules and the pore-size of the gel matrix. Gel electrophoresis is routinely used to separate proteins or nucleic acids; specific protocols have been developed for each type of molecule. Common gel materials are agarose (a polysaccharide) and acrylamide (a 3-carbon amide which is polymerized to form long chains with cross-links between the chains). The pore size of the gel is influenced by the percentage of gel material used and, in the case of acrylamide, the amount of cross-linking.

PAGE (polyacrylamide gel electrophoresis) is used to analyze protein samples.

You will be using gradient acrylamide gels with a continuum of acrylamide concentrations along the length of the gel. Gradient gels are useful because a greater size range of proteins can be effectively separated on a gradient gel compared to a single-percentage acrylamide gel. The gels you will use have a gradient range from 4-20% acrylamide.

Removing Seal:
The gels you will use have a gradient range from 4-20% acrylamide and are contained in a disposable cassette. The solution used to store the gel contains sodium azide. The gel cassettes have a seal over the bottom to prevent leakage when the liquid acrylamide was poured. In order for a current to pass through the gel to move the protein samples, both the top and bottom of the gel need to contact the electrophoresis buffer. Before running the gel, it is necessary to remove the plastic piece sealing the bottom of the cassette:

1. Place the cassette on the bench top with the bottom piece extending beyond the bench.
2. Hold the cassette steady by pressing down on the two plastic pieces that run along the sides of the gel (it is important to anchor both edges);
3. Wiggle the bottom piece until it snaps off.

Marking Well Locations:
In order to facilitate loading of the samples, it is very helpful to mark the locations of the wells. When the comb is removed and buffer fills the wells, it is very difficult to see the wells.

1. Position the cassette so that the taller plate is face up.
2. If the plate is wet, dry it with a Kimwipe, then use a Sharpie marking pen to trace the outline of the wells (it is most important to mark the lower part of the wells.
3. Gently remove the comb.

Buffer is a solution used to maintain proper pH for a procedure.

Preparation Instructions:
Check the concentration of the electrophoresis buffer in the lab. The concentration used to run the gels is "1X". If only the 10X concentrate is available, you will need to prepare a 1/10th dilution (the 10X solution is 10 times more concentrated than the solution required to run the gel). The designations 1X and 10X don't refer to specific buffer contents, but only to the relative concentrations. The buffer we will use contains Tris (= Tris[hydroxymethyl]-aminomethane) as the buffering agent, glycine, and SDS (alternate names: sodium dodecyl sulfate, dodecyl sodium sulfate, and sodium lauryl sulfate). If no 1X solution is available, make up 500 ml of a 1X Tris-glycine-SDS buffer.



Instructions:
With your finger, apply a thin coating of sealant to the gasket on the gel electrophoresis apparatus (informally known as a "gel rig"). The sealant helps to form a better seal between the rig and the gel cassette, minimizing leakage. Clamp the gel cassette into the gel rig with the notched plate facing the upper buffer chamber.


Note: Orient the clamps so that the broad side is facing out. Pour 1X electrode (electrophoresis) buffer into the upper buffer chamber of the gel rig. The sample wells of the gel should be submerged, but the level of buffer should not be higher than the taller plate of the gel cassette (to avoid a "waterfall" of buffer running down the gel plate). Before proceeding further, look very carefully for leaks (buffer running down the plate or a drop in the buffer level of the upper chamber). If no leaks are apparent, fill the lower chamber half to three-quarters full with 1X electrode buffer.


Practice Loading Samples on Gel

Instructions:

1. Place a fresh gel-loading tip on the micropipettor.
2. Take up 5 microliters of practice gel loading solution.
3. Carefully insert the thin tip into a sample well.
4. Eject the sample by slowly and steadily pressing down on the plunger of the pipettor without stopping--this should be one smooth, steady motion. The sample buffer is denser than the electrode buffer that fills the reservoir--this allows the sample to settle into the bottom of the well.

Caution:

* Do not release the plunger before the whole sample is ejected (this will cause mixing of the electrode buffer and the sample buffer and will cause the remaining sample to have less density).
* Do not press the plunger past the first stop position (this will push an air bubble into the well, which may stir up the neatly laid sample). Pull the pipettor completely away from the well before releasing the plunger to avoid sucking up the sample or buffer.

Loading Samples on Gel

Instructions:

1. Place a fresh gel-loading tip on the micropipettor.
2. Take up 5 microliters of sample.
3. Carefully insert the thin tip into a sample well.
4. Eject the sample by slowly and steadily pressing down on the plunger of the pipettor without stopping--this should be one smooth, steady motion. The sample buffer is denser than the electrode buffer that fills the reservoir--this allows the sample to settle into the bottom of the well.

Caution:

* Do not release the plunger before the whole sample is ejected (this will cause mixing of the electrode buffer and the sample buffer and will cause the remaining sample to have less density).
* Do not press the plunger past the first stop position (this will push an air bubble into the well, which may stir up the neatly laid sample). Pull the pipettor completely away from the well before releasing the plunger to avoid sucking up the sample or buffer.

Hint:
Whenever possible, lanes 1 and 10 should be left empty (samples loaded in end lanes sometimes run differently than samples in interior lanes, a phenomenon known as an "edge effect"). It is also helpful to load the samples in an asymmetric pattern, which makes it easier to distinguish the right and left sides of the gel.



Electrophorese Samples


1. Once the samples are loaded, slide the lid onto the apparatus so that the electrical leads are connected to the electrodes (be sure to match the color-coding of the leads and electrodes).



2. Connect the leads to the power supply--black indicates the negative electrode, red the positive electrode.



3. Adjust the power supply as directed by your instructor (you want to run the gel at 100 volts).

Gel is now ready to run!

Caution: Monitor the gel's progress frequently!

* Monitor the progress of the samples as they move into the gel--if the dye begins to move up and out of the wells rather than down and into the gel, turn off the power supply and reverse the electrical leads.
* Sometimes leaking buffer will cause the gel to stop running.



Finished Gel: Remove from Apparatus

Instructions:

After the electrophoresis is finished:

1. Turn off the power.
2. Disconnect the leads.
3. Remove the cover of the gel apparatus.
4. Pour the buffer into the sink (if you don't, you could have a flood when you remove the gel cassette).
5. Remove the gel cassette from the gel rig and blot off excess buffer with a paper towel.
6. Carefully open the gel cassette (you'll need to crack the cassette open using a butter knife-spatulas bend too easily).
7. Coax the gel to lie on one of the plates (the gel may tear if it is handled too roughly-the low percentage gels are especially fragile).
8. Notch one of the upper corners of the gel so that you can keep track of the gel orientation through the staining process--be sure you know which corner is notched relative to the order of loaded wells.


Stain or Develop Gels

You can either transfer the proteins to a membrane from your gel for western blot analysis, or stain the gel directly with coomasie blue stain or other stain.

Here we will be staining with coomasie.

Coomassie Blue


Instructions:
The staining solution contains a dye to visualize the protein bands and alcohol to "fix" the proteins and keep them from diffusing in the gel. (Coomassie Blue [alternate name anazolene], methanol and acetic acid).

Store the Gel(s)

Instructions:
To store the gel after destaining, wrap in plastic wrap, label, and place in the refrigerator. During the next lab, you will use the Gel Documentation system to photograph the gel (to be included in your write-up).

Record your results with the gel documentation system.

[anoopbal]

Thanks for the protocol.

What is the benifit of staining the gel using commasie blue? I am assuming unless you use aan antibody for a specific protein you cannot differentiate your band from other bands.

Thanks

[bdekker]

Hi,
If you need (even) more information, there is a freely available Nature Protocol for SDS-PAGE that you can access from the 'Sample Protocols' of the Nature Protocols website.

[admin]

Thats a decent protocol however if you are looking for more detailed protocol see the protocol SDS-PAGE protocol

[zman]

Thanks ^ it was almost a replica of what we did in the lab! Really useful protocol.

[Satanay]

hello I was wondering if I should load empty wells with sample buffer, my bands become wider when there are empty wells.
thank you

[admin]

Yes, that is a good point Satanay.

Always make sure to fill all empty lanes with sample buffer. You can get weird migrations if even a few lanes are not the same salt/buffer.

[stewdew]

Hello...i'm from Norway and new to this forum, which i now hope to visit frequntly in the future.

I'm in my last year of bachelor in biotechnology, and last week we had an experiment in biochemistry with SDS-PAGE where the sample showed no migration at all. Just some drifting off to the sides, and the teacher did not bother to photograph.
As this was a school experiment we did not run the samples over, and the teacher said it could only be that no power had run through the gel. But a second group shared the gel machine and theire gel migrated very nicly, as did all other groups on different machines.
In the prosess of setting up the machine and gel with buffers we checked for leakage and it all looked fine. Unfortunately as we had a different experiment going on as the gel electroforeses was running, it was stressful and we forgot to check up on the migration, and probable left it running too long, although as said the other group got nice results.

So please if anyone have any suggestions as to what went wrong or why this might have happend please share it with me so i get a better understanding, than a just no power explanation. I'm thinking the machine was broken or something on one side of the gel connections.

Anyways, thanks alot! Stewdew

[admin]

This is a perfect example of (probably) filling the apparatus too high with buffer.
A lab technician explained to me (he knew the process through quite in detail) that any small holes in the apparatus/gel can be quite detrimental to the flow of buffer ions that move your proteins through the gel.

If there is no buffer where there are holes, the ions have no choice but to go through the gel first and then out the bottom.

however, by filling the buffer higher, you are allowing any buffer ions/electrical current to go through the minute holes/defects in the system, minimizing the electrical current where you need it most (through the gel)

and thus slowing migration down a lot... you must have been unlucky and got a bad system, however next time filling the buffer to just above the bottom electrode will prevent this.

cheers