SDS-PAGE
Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis
Equipment & Consumables:
Dedicated workspace - Polyacrylamide can break down into acrylamide, which is a toxic carcinogen. Use masking tape to mark off a section of workbench that is ONLY used for SDS-PAGE.
Set aside a special SDS-PAGE pipette, sacrificed to the acrylamide
10 µl Long-tipped tips make loading easier.
4x SDS-PAGE Loading Buffer. To make 10 ml of 4x stock, Mix the following:
2.5 ml 1 M Tris-HCl pH 6.8
0.5 ml of ddH20
1.0 g SDS
0.8 ml 0.1% Bromophenol Blue
4 ml 100% glycerol
2 ml 14.3 M β-mercaptoethanol (100% stock). You can keep this separate and add later.
Check that it has β-mercaptoethanol added by smell (worst smell ever).
If you do add it, find a way to double or triple contain it to prevent the smell leaking out.
Can be replaced by DTT which smells better
Adjust the final volume to 10 ml with ddH2O
Mix with samples in 1:3 - Buffer:Sample Ratio
SDS-PAGE 10x Running Buffer
95°C Heat Block or Hot Water Bath
Or: PCR Tubes & Thermocycler set to 95°C
Pre-Cast SDS-PAGE Gels appropriate to protein you plan to purify.
e.g. In Open Insulin, our expected protein size varies from 8.9 kDa to 30.1 kDa. Gel+Buffer combos that can resolve these sizes include;
Tris-Glycine Buffer, 4-20%, 12%, 15%
MES Buffer, Bis-Tris 12%
You can cast your own SDS-PAGE gels, but it’s a massive pain.
Plastic Tray larger than gel
Rocking Table
Coomassie Blue Staining Solution
Destain Solution
Optional: Microwave
Kimwipes
Ice in Ice Box
White Light Transilluminator
Or: Smart phone with white screen and a piece of perspex to cover it.
Don’t get acrylamide on your smartphone…
Protocol:
Equip PPE, it is very important during all stages of this protocol.
Write up a load order for your gel, leaving sufficient lanes for the protein standard, as well as each purification step for each sample.
If you have access to a nanodrop, measure A280 for each sample you plan to add to the gel. Use this number to determine the optimum amount of sample to get at even amount of protein in each well.
Between 25 and 50 µg per lane usually works out well. For example if your sample is 5mg/mL that’s equal to 5µg/µL so for a 40µg lane you would need 8µL of it.
Label (or number based on well) microcentrifuge or PCR tubes for each sample and then add this quantity of sample to your fresh tubes. Note the amount of sample added to each on your load order.
Mix each of your samples in a 1:3 ratio with 4x SDS-PAGE Loading Buffer by pipetting up and down in the tube. e.g. Add 2 µl of SDS-PAGE Loading buffer to the 6 µl of Eluted protein above for a total of 8 µl.
Consider adding up to 50% more sample and loading buffer than you actually need to mitigate loss during the next step.
Heat samples to 95°C for 10 minutes in the Heat Block/Hot Water Bath or Thermal cycler. Note that the protein standard should not be treated this way, it is ready to load.
Alternatively you can do 100°C for 3 minutes, but this may cause greater loss of sample volume.
Take a pre-cast gel cassette, remove the packaging and take off the strip from the bottom of the gel (Don't leave this on or nothing will run!). Remove the comb with the utmost care not to damage the wells.
Try to hold the gel on either side, pinch the comb from either side and then slide it out evenly. Pulling the comb out unevenly can ruin the entire gel!
Slide the gel into the clamp and affix it in place by clipping up the sides. If you're only running one gel, you will need to use the perspex 'buffer dam' on the other side of the gel holster core. If you're running two gels, affix one on either side.
The mechanism and shape of your SDS-PAGE equipment will affect this step quite a bit. This protocol is for the Biorad apparatus. Adjust your protocol for your specific piece of equipment, googling as necessary.
Fill the inner buffer chamber of each core right to the top with ~200 ml of 1x Running buffer.
If the buffer runs out of the bottom, don't proceed. It's not going to work, take it apart and put it together better / with unbroken parts. If it holds the buffer without much running out, you can proceed.
Note: "Each core" is used because some biorad chambers have 2 cores for a total of 4 gels. If using one of these, you will need more buffer outside of the gels (marked on chamber).
Fill the outer buffer chamber to the indicator mark for 2 gels (550ml) or 4 gels (800ml) if using multiple gels in the large buffer chamber.
If you are running your gels at >200V, you should also use 800ml in outer chamber.
Using your load order as a guide, add your protein standard and each of your samples to the wells of the SDS Page gel.
Loading for these gels is notoriously difficult. Long-tipped tips can help here, as can clear markings of the well locations.
If sample prep went well, the sample should sink to the bottom of the well and sit relatively evenly.
Minimise air bubbles, especially within the tip prior to loading.
Attach the lid to the top of the chamber, ensuring the correct electrode plugs are aligning properly to the electrodes.
There is a clearly marked +ve (red) and -ve (black) side for both chamber and lid.
Plug the electrodes into the power pack and then run your sample at 200V for 31-39 minutes.
This is simple a starting time and voltage. It may be necessary to vary this to obtain the best possible resolution.
Check the instructions that come with the gel to ensure you're using the right voltage for your specific gel type. It varies!
Turn off the power supply and disconnect the electrical leads. Remove the lid.
Remove the gel holster core containing your gel and pour the buffer from the central chamber back into the buffer chamber. Remove the wings holding the gel in place and place the gel cassette onto some kimwipes or paper towel, with the shorter plate on top.
Slide the cracker in at each point of the cassette indicated by the arrows and twist up (crack!) while being careful not to disturb the gel between the glass plates. After doing this at each point, you should be able to lift off the front glass plate, exposing the gel.
Using your jet bottle, spray water to push the gel off the bottom plate and into a plastic tray with enough room for it to move.
The gel is incredibly easy to tear, try avoid touching it unless absolutely necessary. Keep it wet and move it around using water pressure whenever possible.
Pour excess liquid out of plastic container while gently holding the gel in place.
Add enough Coomassie blue staining solution to cover the gel by 1.5 cm. Incubate the gel in the stain on a rocking table for 1 hour.
Optional: Microwave the tray on high power for 40-60 seconds. The staining solution boil will boil so include some kind of cover over the tray. Allow to cool on the rocking table for 10 minutes.
This protocol is faster, but may result in less clearly distinguishable bands.
Pour off the used Coomassie stain into liquid waste.
It can be reused 2-3 times if you can filter purify it.
Rinse the gel in the container 2-3 times with dH2O to remove as much Coomassie stain as you can from the gel & container.
Add fresh Destain solution to cover the gel by ~2cm. Tie ~4 kimwipes into knots and lay them around the outside of the staining chamber to adsorb excess stain.
Try to avoid laying the kimwipes over the gel itself, this will cause uneven destaining.
Incubate on the rocker for at least 1 hour or preferably overnight.
Optional: Microwave the tray on high power for 40-60 seconds. The destain solution will boil so include some kind of cover over the tray. Allow to cool on the rocking table for 10 minutes.
This protocol is faster, but may result in less clearly distinguishable bands.
Discard the kimwipes and replace with fresh kimwipes. Destaining step can be repeated/left longer if too much background stain remains.
Illuminate and photograph your gel using a white light transilluminator.
Need something clear to lay the gel on and a white background.
Don't overthink it. Perspex and a phone screen is okay. Try build something for long term, but no dedicated equipment yet.
Note: If protein continuously fails to appear, consider also collecting supernatant from each wash step during the purification protocol to see if your protein is being eluted too early or failing to bind.
Acknowledgements:
Coleman Protocols 2017 & 2019 http://coleman-lab.org/