Could that be a massive protein I see? We can separate proteins by size by sending them traveling through an SDS-PAGE gel, but in order to find our protein treasure we need to send in some treasure hunters. And the one we use most often is a dye called Coomassie Brilliant Blue (CBB), either in the classical way or in the form of faster colloidal stains (often advertised as some trademarked version of “instant blue”). 

note: this post is refurbished from September 2019, about 300 SDS-PAGEs ago (literally, not exaggerating – today I ran my 732nd, 733rd, & 734th!)

SDS-PAGE (Sodium Dodecyl Sulfate – PolyAcrylamide Gel Electrophoresis) is a technique we use to separate proteins in a mixture by size (well, length) using electricity to send the proteins through a gel mesh. The goal here isn’t to purify the proteins, instead the goal is just to see what’s there. But protein’s are invisible to the naked eye, so we need some sort of dye in order to protein-spy.

note: the blue you see while the gel is running is just a tracker dye in the sample loading buffer which helps you know when to turn off the electricity and start the staining. 

You can think of the SDS-PAGE gel’s matrix as maze with protein “treasure” hidden throughout. Unlike the proteins, with moved straight down the gel because the electric field was compelling them that way, the dyeing step happens after you turn off the electricity and transfer your gel from its plate sandwich to a staining box. Here, there’s no directionality to movement of things, so the dye will move wherever the heck it wants! (though the protein will be stuck in place through chemical-induced clumping as we’ll see). 

The dye travels randomly through the gel maze by diffusion (the molecules move around randomly, ricocheting off the things they run into with the NET RESULT that they move from areas of high concentration to areas of low concentration). When it finds protein treasure, it latches on. And since the dye’s blue it tells us where the protein is.

But it’s really hard to find treasure that’s trying to escape! So we need to get the treasure to stay put – the FIXATION step uses an alcohol and/or acid to make the protein to precipitate (clump up) so it gets stuck in place so our treasure-hunting dye can find it. We also have to watch out for overly-eager treasure hunters that latch onto “fool’s gold” (stain the gel itself) causing high background. This leads to an overall treasure-hunting scheme of:

  1. GEL ELECTROPHORESIS – separate proteins by size by unfolding them, coating them in negatively-charged detergent (SDS) & using that negative charge to motivate them to travel through a polyacrylamide gel mesh towards a positive charge. The bigger (longer) proteins get tangled up more, so they travel slower and have progressed less (so higher up on gel) when you turn off the power. More here:
  2. WASH – remove free SDS, etc. 
  3. FIX – trap the treasure – the gel has to allow proteins to move when you want them to, but not too easily – Ideally they’d only move when power’s on, but molecules like to move and if they can they will – so (especially the small ones) can start wandering off even when the power’s off). To prevent that wandering, you add an alcohol and/or acid to get them to clump and get stuck in place. 
  4. STAIN – send in the treasure hunters – stick the gel in a bath of dye – the dye enters, latches onto the protein and gets stuck too (this step’s often combined w/the fixing step)
  5. DESTAIN – call off the hunt – get the treasure-less treasure hunters to leave so you can better see where the treasure-full ones are (some rapid stains have low backgrounds & you just destain in water, letting the dye diffuse out, but some methods use more complex destaining)

As I alluded to, there are lots of different formulations including “Classic Coomassie” which has really eager treasure-hunters that can find tiny amounts of treasure but are also fool’s-gold-happy – it’s super sensitive and eventually gives you nice crisp bands, but takes a lot of destaining to reveal them. Alternative “Colloidal Coomassie” recipes are becoming more and more common because they’re faster & more eco-friendly – these forms keep groups of treasure-hunters hanging out outside the gel and gradually send them in until all the treasure’s found and bound.

Let’s take a closer look at our treasure hunter, Coomassie Brilliant Blue (CBB). Most scientific reagents have boring (though descriptive) names, so you might wonder how Coomassie Brilliant Blue (CBB) got its name. If you’re kinda nerdy, you might look it up. And if you’re even nerdier like I am, (embrace it!) you might then tell other people about it!

“Coomassie” is actually a trademark name – owned by a company that no longer even makes it – & a town name (modern-day Kumasi, Ghana). CBB wasn’t discovered or produced there, nope – a British company thought naming their product after the capital of the Ashanti empire they recently conquered would be good business strategy. That makes me mad, so I’m going to call it CBB most of the time. 

“Coomassie” was initially used to market a wide range of wool dyes, with CBB first made in 1913 & first used to stain proteins in 1960s. Dye-ing to know more about the dye itself? It has a pinwheel-like chemical structure and is characterized as a triphenylmethane dye. Unlike the common name a company decided to give a product, this is an example of a functional name – it describes the chemical makeup – in this case three (tri) phenylmethane groups – phenyl is a type of resonance-stabilized ring (a ring or atoms that share electrons in a delocalized fashion that makes them good at absorbing light and thus making things look colored) – and methyl is a -CH₃ group.

There are 2 main forms of this “triphenylmethane” dye, R-250 & G-250. Both are blue, but R’s more “reddish” & G’s more “greenish” (although the color depends on the pH and whether and what it’s bound to which is why it can be used to measure protein concentrations in something called a “Bradford assay ). “250” was originally a purity/strength indicator. R-250 is more sensitive , but G-250 can be made into forms that produce lower background, with faster protocols.

We get the G-250-based “quick stain” we use most of the time pre-made, but we make our own “Classic” R-250 stain. It’s a really simple recipe –  only 4 ingredients: water, acetic acid (AcOH), methanol (MeOH), & CBB – but it takes a while to prepare… more here:

But none of that really matters if the dye’s not where you want it – and only where you want it. Where we want it is stuck to our protein (which is itself stuck in our gel). And where we don’t want it is anywhere in the gel there isn’t any protein. So how does it stick to our protein but not the gel? The bumbling biochemist’s here to tell!

CBB has sulfuric acid groups that can be negative or neutral depending on pH. Under the conditions of the staining solution it has overall ➖charge (anionic), so it binds (reversibly) to ➕-charged parts of proteins (basic amino acids like Arg, Lys, & His) through electrostatic interactions (opposites attracting). note: those side chains aren’t always positive, but we stain the gel in acidic conditions, where they’re more likely to be – more here:

CBB also binds to non-charged protein parts (especially the ring-y (aka aromatic) amino acids like Phe, Tyr, & Trp) “generically” through “Van der waals” interactions, which involve shifting around of electrons when molecules get close together.  These interactions are individually weak but they add up (they’re what allow geckos to walk up walls!). Proteins with unusually high proportions of ring-y amino acids tend to stain better. An example is BSA (bovine serum albumin), which recruits twice as many treasure hunters per weight of protein than your average protein.  

Speaking of weight, different proteins have different weights because they have different #s of amino acid letters. We commonly talk about protein weights in terms of “molecular weight (MW)” and units of “kiloDaltons” (kDa). BSA’s molecular weight is 66.5 kDa, meaning that 1 mole (6×10^23) of BSA molecules would weigh 66.5 kg. More on moles here:

But the key thing here is that bigger proteins have “more to love” in CBB’s eyes – they offer more binding sites and thus will more BSA per protein molecule than a smaller protein. So your bands would look darker for bigger proteins than smaller proteins if you ran the same # of protein molecules of each. 

CBB’s protein-binding ability also makes it a good wool dye because wool’s chock full of a protein called keratin. And that keratin-binding ability which makes it good at dyeing wool also makes it good at dying your skin – so wear gloves & avoid splashing. CBB can also bind SDS (that detergent we used earlier to solubilize and negatively-charge the proteins) which can mess up results. So you want to wash your gel in water before staining to remove the SDS.

The classical CBB method goes something like this (apologies for scribbled-note-like formatting)…

  1. water wash -> get buffer components to diffuse out of gel -> start with a “clean slate”
  2. fixation/stain -> immerse gel in a staining solution (CBB + methanol (MeOH) + acetic acid (AcOH)) (some more eco-friendly methods use ethanol)
    • we dissolve CBB in a MeOH/AcOH solution to combine the fixation & staining steps. note: It takes a really, really long time for the CBB powder to dissolve, so let it stir for several hours on a magnetic stir plate and then filter through a coffee filter to remove any undissolved stuff 
    • fixation: protein precipitates so is trapped in the gel matrix
    • stain: dye diffuses into gel
      • when dye meets protein -> dye binds -> gets trapped because the protein’s trapped
      • where there’s not any protein, the dye still fills the pores of the gel, but it’s not trapped (it can flow in and out but at this point its flowing mainly in because the dye concentration ([dye]is higher in the stain than in the gel).
      • When [dye outside gel] = [dye inside gel] you’ve reached “equilibrium.” Equilibrium isn’t a stop point – it’s actually a dynamic process (the individual dye molecules are still moving around but the rate of molecules entering the pores = rate exiting the pores, so you can’t tell anything’s changing.
  3. destain -> swap out the stain solution with a destain solution (MeOH + AcOH but NO CBB) -> remove the non-protein-bound CBB
    1. the stain can’t go down the drain – instead we collect it in hazardous waste containers that our environmental health & safety (EH & S) workers dispose of properly
    2. at the start, there’s high CBB in gel, NO CBB in bath -> CBB diffuses out of gel into bath
    3. this makes CBB concentration in bath increase -> less of a difference between gel & bath -> CBB diffuses more slowly

As you can hopefully see from the pics, Classic CBB initially stains your whole gel blue so you can’t see the bands until you destain it and de-blue what’s not supposed to be blue. This is because in order to find tiny treasure you unleash a ton of treasure hunters that race in from the dye bath, where there’s a high concentration of dye, to the pores of the gel where there’s more room – and hopefully protein! But they’ll keep snooping around even if there’s no protein left to find because you have high concentrations of free dye inside and outside the gel.

But Colloidal CBB doesn’t require as much destaining (though it helps make the bands crisper) because, instead of sending in a ton of treasure hunters you then have to kick out, it has groups of hunters hang out outside the gel and only go in “as needed” 

A colloid is a solution where instead of having individual molecules spread throughout, you have “clumps” of those molecules, but still evenly spread out like in a “normal” solution. And those clumps really are “dissolved” (they don’t come “undone” when you let the solution sit).

The colloids are too big to enter the gel so you don’t “overstain.” BUT the FREE molecules can come & go from the gel as they please – and they do, just as before, except now they’re at a lower concentration. So how do you get enough to stain the protein? No problem –  the free CBB population can constantly get replenished from colloidal “stock rooms” (you have a sort of equilibrium between free & colloidal).

The result? Instead of flooding the gel with dye (like the “classical” method) you supply a steady, mild flow of CBB molecules – enough to quench the proteins’ thirst, but not so much that it can stick to the gel.

Thus, you can see the bands even without destaining. It also helps that, although we think of CBB as blue, it’s color depends on its charge. At low pH, CBB G-250’s brownish, but when it binds proteins, it gets stabilized in blue state so you can tell background brown from treasure blue.

At least, the stain we used to get was more brown-y. That stain’s been on back-order for months. And months. And months. So we’ve been experimenting with a few other formulations but they seem to start bluer and give higher background. 

Speaking of color, a great thing about coomassie stains is that, unlike the DNA gels that use fluorescent dyes we have to stick on a UV tray, CBB is “colorimetric” – the readout is color, so the results are literally right there to see, no special equipment required (unless you count the eye, which is a pretty spectacular tool!) (and the light tray helps too 🙂 ).

Another colorimetric protein stain is silver staining, which is more sensitive, but also more finicky. 

There are also fluorescent protein stains. CBB & silver stain are both “all protein stains” – they stain all proteins (though sometimes not equally well). There are also fluorescent all protein stains. But the really cool fluorescent stains only stain specifically-modified proteins like glycoproteins (which have sugar chains added) or phosphoproteins (which have phosphate groups added) 

some sources on CBB history and use:

more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉


#scicomm #biochemistry #molecularbiology #biology #sciencelife #science #realtimechem

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