SDS-PAGE
Coomassie Brilliant Blue binds protein through ionic and hydrophobic interactions and is the default stain: about 60 ng per band for R-250 in methanol/acetic acid, roughly 15 ng for colloidal G-250, and a response that stays near-linear over about two orders of magnitude.
Our ruleSimulations prepare you for the bench but don't replace hands-on practice.
Coomassie stain
Coomassie Brilliant Blue binds protein through ionic and hydrophobic interactions and is the default stain: about 60 ng per band for R-250 in methanol/acetic acid, roughly 15 ng for colloidal G-250, and a response that stays near-linear over about two orders of magnitude.
How it differs from the standard method
- Near-linear response makes it the stain to use when band shares are part of the answer.
- Colloidal G-250 needs no destaining step and is more sensitive, but takes several hours to develop.
- Fully compatible with cutting bands out for mass spectrometry.
1Why a gel can measure a mass
Proteins differ in charge as much as they differ in size, so an untreated protein mixture separates on a gel by a tangle of charge, shape and mass that no single ruler can read. Sodium dodecyl sulfate removes two of those three variables.
Make the band thin
A large-pore 4% gel at pH 6.8 sits above the resolving gel. Glycine is barely charged there, so chloride runs ahead and glycine behind, and the proteins are swept into a disc microns thick between them.
Sieve by size
At pH 8.8 glycine becomes fully charged and overtakes the proteins. The stack collapses and each protein migrates at a rate set by how it threads the mesh.
Set the window
7.5% resolves roughly 35–350 kDa, 12% about 10–120 kDa, 15% about 5–70 kDa. A gradient gel steepens as the proteins descend and covers both ends.
See the bands
Coomassie R-250 detects about 60 ng per band and responds roughly linearly; silver detects a few ng but saturates fast.
2What the readout physically is
The number you report is not a mass. It is a relative migration, how far a band travelled as a fraction of how far the dye front travelled, converted into a mass by comparing it with marker proteins run in the same gel.
3The four ways it lies
- Glycosylation
- Carbohydrate adds hydrodynamic bulk and binds almost no SDS, so a glycoprotein typically runs 10–30% heavy, often as a fuzzy band because the glycan is heterogeneous. Treating with PNGase F and watching the band collapse is the test.
- Extremes of charge
- Very basic proteins such as histones bind less SDS per residue than average and migrate slowly, reading high. Very hydrophobic and multi-pass membrane proteins bind more and often read low, sometimes by 20–30%.
- Reduction
- A reducing agent breaks inter-chain disulphides before the gel. A covalently linked dimer and two separate monomers then give exactly the same band, so a reduced lane alone cannot tell them apart.
- Incomplete denaturation
- SDS at room temperature does not unfold a stably folded protein. Part of the sample stays compact, runs ahead of its true size and looks like a smaller species: a common reason for an extra band.
4Loading, and what overloading does
A Coomassie band needs roughly 60 ng to be seen and distorts above roughly 3 µg. Between those two numbers lies the whole useful range of the gel, about 50-fold, and it has to accommodate both the protein you care about and the contaminant you are trying to spot.
5Leaving the SDS out
Native PAGE runs the same acrylamide system without SDS, without a reducing agent and without heat. Migration then depends on charge, shape and size together, so the apparent mass means little, but the complex survives the gel, which is the one thing SDS-PAGE can never offer.
Key terms
Practise these as flashcards →Distance a band travelled divided by the distance the dye front travelled. The quantity a gel actually measures; masses are read from it against a marker.
Total acrylamide concentration of the resolving gel. It sets the pore size and so the range of masses the gel resolves.
The discontinuous Tris-glycine buffer system with a stacking gel at pH 6.8 over a resolving gel at pH 8.8, still the standard for protein gels.
Concentration of the sample into a very thin disc at the gel interface before separation begins. It is what makes a sharp band possible.
Whether DTT or 2-mercaptoethanol was added. Only the comparison of the two lanes identifies a disulphide-linked species.
The mass a band reads against the marker, as opposed to the mass calculated from the sequence. The difference between them is often the interesting result.
A resolving gel whose acrylamide percentage increases down the gel, widening the range of masses resolved in one run.
When to use it
- Checking the purity and apparent size of a protein preparation at any stage of a purification.
- Finding out whether a high-mass species is a disulphide-linked oligomer, by running reduced and non-reduced samples side by side.
- Separating a complex sample before transferring it to a membrane for a western blot, or before cutting bands out for mass spectrometry.
- Asking whether a protein is modified: a shift away from the sequence mass is the first evidence of glycosylation or a large covalent adduct.
Compared with alternatives
Adds antibody detection after the gel; use it when you need to identify one protein in a complex sample rather than see everything that is there.
Separates by isoelectric point first, so it resolves proteoforms that share a mass and are a single band on SDS-PAGE.
Gives an accurate mass and an identification rather than an apparent one; use it when the gel's answer is ambiguous or when you need to know what a band is.
Measures the mass of a complex in solution without a matrix, so it answers the oligomeric-state question that native PAGE only hints at.
Tells you how much protein you have before you decide what to load; a gel is a poor way to measure total concentration.
Sub-techniques
Choosing one here switches the whole page, including the protocol and the lab, to it.
Coomassie Brilliant Blue binds protein through ionic and hydrophobic interactions and is the default stain: about 60 ng per band for R-250 in methanol/acetic acid, roughly 15 ng for colloidal G-250, and a response that stays near-linear over about two orders of magnitude.
- Near-linear response makes it the stain to use when band shares are part of the answer.
- Colloidal G-250 needs no destaining step and is more sensitive, but takes several hours to develop.
- Fully compatible with cutting bands out for mass spectrometry.