Silver staining is kinda like being a talent scout & agent for D-list celebrities. You want to find them & make them shine. With a silver stain we can make tiny amounts of protein shine – literally! All it takes is a coat of silver (on the protein not the box….) SILVER STAINING is a sensitive all-protein stain that lets you find invisible proteins trapped in a gel. It’s not as popular as Coomassie, but when it comes to tiny amounts of protein, silver staining does the job more well (pictures English teachers cringing – sorry!)
When I say sensitive, I mean it. It’s ~50X as sensitive as coomassie-based stains, able to detect as tiny an amount as 0.25 ng of protein! (that’s 25-quadrillionths of a gram).
Note: silver stain can also be used to stain for nucleic acids & in some histology stuff – I’m focusing on staining protein gels, but the same principles apply for the most part note 2: in the video and in-depth discussion I discuss Bio-Rad’s Silver Stain Plus Kit because that’s what I use (not a paid endorsement) but I talk about what’s inside them so you can see if the method you use has similar components.
I revamped & pieced together some silver staining posts that I went into more detail starting starting here: https://bit.ly/2szMg4k
So, how do you find a tiny amount of protein? First you need to get it out of the shadow of all the “big stars” – you want to spread out the proteins, which you can do using one of the most common techniques in biochemistry, especially for protein biochemists -the SDS-PAGE. It’s a way to separate proteins by size using electricity to send them swimming through a gel mesh, which slows down bigger ones more so when you turn off the power they’ll not have traveled as far. Lots more on it here http://bit.ly/sdspageruler
This spreads them out, but they’re still invisible (even the A-listers). Enter the talent scouts – use a protein stain to find where all the actors are. Most of the time we use a stain that binds to all proteins & the most common one is Coomassie Brilliant Blue (CBB) staining, which we went over in detail starting here. It binds non-specifically to proteins and makes them look blue. Much more on it here: http://bit.ly/cbbgelstaining
But this talent scout only finds the low-hanging fruit – it isn’t the most sensitive stain so it won’t detect proteins that are present in really small amounts. The silver stain is really sensitive (sometimes too sensitive as we’ll see…) but it’s more complex – pun not originally intended, but I’ll take it!
Most of the time I use CBB and we even have an instant stain version we use – 1 bottle, stick it on & see your proteins – at least the ones there’s a lot of… http://bit.ly/cbbgelstaining
In contrast (another initially unintended pun…) when silver staining, there are lots of ingredients, bottles, & solutions. BUT there are really only 4 “final” solutions that you bathe the gel in (at least in *this* protocol… there are probably even more variations on silver stain protocols than there are on Coomassie protocols, which is definitely saying something… )
this protocol uses:
- stop solution
Regardless of protocol, the basic steps are:
- fixation: lock the proteins in place & remove interfering compounds
- sensitization: increase sensitivity & contrast
- silver impregnation: add silver (in soluble, ionic, form)
- image development: build up solid silver metal image
- stop: stop development before excessive background forms
I’ll take you through the steps in more detail, but first here’s the basic premise of the silver stain. The stain solution contains SOLUBLE silver ions (charged molecules). You can’t see them, but they’re there, and they bind to protein in the gel → they get converted to solid silver metal (NOT soluble) → initially you still can’t see it (there’s a “latent image”) → over time it builds up to the point where → you see bands showing the location of protein.
And then you stop it before it starts over-staining the background! Or else you’ll stop seeing bands because your whole gel will just be black…
That’s the overview – now the deets…
Going back to our talent scout analogy, we first need to scout out the talent – get the dissolved (& invisible) silver ions (Ag⁺) to bind our protein. And then get it to shine -> we’ll use formaldehyde to REDUCE it to neutral, solid (& visible) silver metal
2 Ag⁺ + formaldehyde + 3OH⁻ → 2 Ag + formate + 2H₂O
This is a type of reduction/oxidation (REDOX) reaction, which you can review here: https://bit.ly/redoxbiochem
The key thing to remember is OIL RIG: Oxidation Is Loss (of electrons, e⁻) and Reduction Is Gain (of e⁻)
Different variations in protocols try to balance sensitivity (ability to detect small amounts of protein) & contrast (stain protein but not the gel).
But before we send in those talent scouts, we need to FIX the gel. No, I’m not talking about piecing together the gel you accidentally tore (though I’ve certainly been there!) Instead I’m talking about getting the proteins stuck. It’s impossible to find all the actors at their homes if they’re running around the country, and similarly, it’s impossible to find all the proteins at their finish locations if they’re diffusing around the gel (moving randomly with the net effect of going from where they’re most concentrated (in this case the place they are in the gel when we turn off the power) to the place they’re least concentrated (so out in that bath you put them in…).
The gel mesh helps make diffusion harder, but not impossible. So we want to really “freeze” our proteins in place, which we can do with the FIXATION solution (this is just like is done in some classic Coomassie staining protocols). The fixation solution contains methanol (MeOH) & acetic acid (AcOH) which “precipitate” the proteins. Basically they cause the proteins to come out of solution and form big clumps (aggregates) which are too big to escape the gel’s pores. So they stay put.
After fixing our gel, we WASH it with really pure water* to push the fixative out of gel pores (don’t worry, the proteins are safe because they’re locked in there now). In the washes, the fixative leaves the pores and the pores fill with pure water instead. This is important for a couple reasons.
- Silver-staining is pH-dependent, so we need to remove the AcOH & tightly control the pH
- We need to wash any other molecules that could interfere with the staining process – SDS, buffering chemicals, etc.
The wash is a diffusion-driven process (read: really slow…) – it relies on a concentration gradient between the gel & the bath, so you have to replace the water with fresh water partway through to maintain the gradient.
*We want to remove impurities so it’s important we’re not just adding new one! Therefore, we need to use water that’s super pure (like Milli-Q water). Typical tap water contains dissolved minerals such as table salt (sodium chloride, NaCl). When they dissolve in water, they split into ions (charged molecules), Na⁺ & Cl⁻. These ions can interact with silver ions (Ag⁺) and cause it to precipitate (come out of solution). This lowers the sensitivity (because there’s less Ag⁺ available to bind proteins) & lowers the contrast (increased background staining because Cl⁻ molecules are spread throughout the gel). Water can can also have uncharged particulates. And we need to get rid of all these, which the Milli-Q filter does.
Once it’s fixed & washed, it’s on to the STAINING! Send in the talent scouts! Our talent scouts are
- a SILVER amine complex, [Ag(NH₃)₂]⁺, bound to a CARRIER molecule, tungstosilicic acid (TSA)
- a BASE, carbonate (CO₃²⁻).
And we want to make sure that these silver scouts find the protein BEFORE the REDUCING AGENT, formaldehyde, makes it shine – we’re looking for D-listers, not Z-listers!
Once inside the gel TSA-Ag⁺-NH₃ gives up Ag⁺, which is more attracted to the protein than to TSA. Ag⁺ can bind different parts of proteins, most “obviously” negatively-charged side chains on glutamate & aspartate). In the TSA complex, Ag⁺ is more reactive, but once it binds to the protein, it becomes more stable & less reactive. BUT it’s no longer “shielded” by TSA, so it’s easier for formaldehyde to find & reduce it than when it’s in the TSA complex. This is good or else we’d get even worse background!
As the name suggests, formaldehyde is an aldehyde. ALDEHYDES (a carbonyl (C=O) bound to a carbon group (“R”) or hydrogen (H) on 1 side & H on other side can act as REDUCING AGENTS. This means that they can donate e⁻ to OXIDIZING AGENTS like metal cations (positively-charged molecules), getting oxidized in process.
When an aldehyde donates e⁻ to dissolved silver ions (Ag⁺), the Ag⁺ becomes reduced and without the charge (which water loves to hang out with) it’s INSOLUBLE. It therefore “plates out” as solid silver metal.
We need the silver ions (Ag⁺) to find the proteins in our SDS-PAGE gel BEFORE the activated formaldehyde finds & reduces it, converting it from a soluble ion into an insoluble metal we can see. Unlike with Coomasssie staining where it’s ok if you “forget about it” and leave it in the stain for a while, if you forget about a silver stain, you can “forget about” seeing your protein!
As long as there are plenty of reactants & conditions are basic (as in alkaline), silver will continue to deposit. And once a little does, a lot does, because it’s autocatalytic (silver speeds up its own reduction because it offers up an inviting surface for other Ag to join). If you aren’t really careful, you can over-stain. As you admire your protein bands, the whole gel will get coated, hiding them!
And this happens REALLY QUICKLY once it gets going!
note of warning: Another consequence of the autocatalytic nature, is that if the first “trendsetting” reduction occurs where we don’t want it (such as gunk in the gel box that got stuck to the outside of gel, you can quickly develop artifacts (basically false signals). So it’s super duper important to make sure your gel boxes are super duper clean. Not just the quick rinse out you normally do with gel boxes. Instead, I clean with nitric acid to dissolve any residual metal, turning the solid silver into AgNO₃.
So, how do we stop it? It isn’t basic – it’s acidic! Water (H₂O) can split into hydroxide (OH⁻) & H⁺ (a proton), but the H⁺ doesn’t really exist because it quickly latches onto an H₂O to form H₃O⁺ (hydronium ion)
- When there’s more H⁺ (H₃O⁺) than OH⁻, we call a solution acidic
- When there’s more OH⁻ than H⁺ (H₃O⁺), we call a solution basic/alkaline
By looking at our reaction
2 Ag⁺ + formaldehyde + 3 OH⁻ → 2 Ag + formate + 2 H₂O
We can see that we need OH⁻. That is, it relies on BASIC (ALKALINE) conditions. note that formaldehyde goes “all the way” to formate in these conditions because OH⁻ can deprotonate (remove an H⁺) from formic acid
We keep a close eye on the gel as it develops &, when we decide our bands are strong enough, we STOP the reaction by adding ACETIC ACID. This acts as an acid (donates a H⁺) to neutralize the OH⁻, removing this reactant & stopping the staining! Hopefully not too late!
We only want to see where our protein’s located. And the reason silver lets us do this is, well, there are actually multiple reasons – multiple ways silver can bind to the protein. Ag⁺ is hydrophilic (water-loved) because it’s positively-charged and can interact with water which is polar (has partially positive (δ⁺) & partially negative (δ⁻) parts).
Proteins also have areas of full or partial negative charge which, Ag⁺ is happy to hang with. These protein regions include:
- negatively-charged amino acid side chains. These are the parts that stick out from the “generic” protein backbone & make each of the 20 amino acids unique. The “acidic” amino acids, glutamate and aspartate will be deprotonated & negatively-charged under the basic conditions of the reaction (as may some others depending on how low the pH is)
- “naturally but neutrally” electron rich parts
- parts that, while not fully charged, are δ⁻, or can be “induced” to be δ⁻ by Ag⁺. Basically, Ag⁺ can draw weakly held e⁻ towards it, creating temporary dipoles (charge separation) in normally nonpolar parts. This allows for weak electrostatic interactions.
So silver ions can bind protein. Then formaldehyde can reduce the silver ions to solid silver, which is solid. Why’s it solid? Well, we talked about how the silver ions are hydrophilic. The same canNOT be said about solid silver. When formaldehyde reduces Ag⁺, it’s adding an electron, which, being negatively-charged, cancels out that positive charge that was getting water to chill with it. So it cozies up with other neutral silver atoms instead. Water avoids the whole lot and, voila! Solid silver precipitate.
But, especially because of that whole autocatalytic thing, we want that initial reduction to occur ON THE PROTEIN & we need to make sure the nearby Ag⁺ molecules are VERY nearby -not just “in the gel.” How do we do this? There are LOTS of “mix-&-match” approaches to “sensitization” (>60 published protocols in the ‘80s alone).
Some methods control WHERE the trendsetter Ag⁺ binds BEFORE you “activate” the reduction
- make PROTEINS *more* attractive to Ag⁺ by pre-binding other compounds to the protein that also bind Ag⁺ . This is referred to as SENSITIZATION and there are a lot of different compounds used for this including sodium thiosulfate
- this also allows for AMPLIFICATION because it increases the number of Ag⁺ binding sites on your protein
These sensitization strategies can be used alone or together with methods that make the BACKGROUND *less* attractive to Ag⁺, improving the contrast between protein signal & background noise
- thoroughly wash out electrophoresis buffers
- add chemicals to sequester un-protein-bound Ag⁺ co it can’t react
Other methods speed up the start to promote the initial reduction by pre-binding reducing agents (such as glutaraldehyde) to protein so they’re right there when you’re ready to initiate the reduction.
And still others slow down the continuation of silver plating to give protein time to loosen it’s grip on bound Ag⁺ so it can be reduced on-site. This can be achieved by using mild reducing conditions.
Now some more details on the various steps. Starting with preparing the mixtures. In the silver staining kit I use, there are a number of tubes with solutions you have to mix together in the right order at the right time.
And you also have a few “not included” things – to prepare the fixative you mix the methanol, acetic acid, water, and “fixative enhancer concentrate) which has glycerol which can keep your gel from shrinking too much and somehow reduce background.
After the fixation you do a couple of water washes (you need a couple since diffusion just goes until an equilibrium is reached inside & outside the gel, and the stuff that leaves the gel in the wash is then outside the gel…)
While you wait, (while the gel is soaking in the water) you can prepare your stain solution. This is a good time, not just because it saves you from boredom, but because you don’t want to prepare your stain until right before you’re ready to use it. This will help prevent unwanted “side reactions” from occurring such as formaldehyde polymerization (formaldehyde has a tendency to link up into chains) or the precipitation of silver ions (Ag⁺) by carbonate.
I’ll make the stain in 2 parts & then add it to the gel once it’s fully made.
- solution 1) “Silver Complex Solution” = silver nitrate (AgNO₃) & ammonium nitrate (NH₄NO₃)
- solution 2) “Reduction Moderator Solution” = tungstosilicic acid (TSA)
- solution 3) “Image Development Reagent” = formaldehyde (HCHO)
part 2: “Development Accelerator Solution” = sodium carbonate (Na₂CO₃)
- note: make this fresh by dissolving the powder in water (and the really pure stuff, remember!)
so why all this complex ordering?
Let’s start with solution 1, our “Silver Complex Solution.” It contains a mix of silver nitrate (AgNO₃) & ammonium nitrate (NH₄NO₃) dissolved in water. Because they’re dissolved in water, there’s a mix of ions swimming around in there (Ag⁺, NH₄⁺, & NO₃⁻)
Right now, the NH₄⁺ wouldn’t want to hang out with Ag⁺, because they’re both positively-charged. BUT later, when we add another chemical to lower the pH, the NH₄⁺ will deprotonate to give you NH₃, which is neutral and can form a complex with the Ag⁺ ions
I use a “diamine” stain: “di-” = 2 + “-amine” for “ammonia.” Each Ag⁺ molecule will form a complex with 2 ammonium particles. And these complexes keep the free Ag⁺ levels low so you get a lower background. (You still need some free silver so that it will be able to get into the gel, but you don’t want a ton). Some methods use silver nitrate without the ammonium but with these stains, you have a much higher free Ag⁺ concentration, which can give you a much higher background staining if you don’t use some of those “enhancement” tricks.
First we mix solutions 1 & 2. Conditions at this point are acidic (low pH), so ammonia & TSA still mostly have their protons (H⁺) so they’re not very reactive with Ag⁺
Next we add solution 3. Once again, the acidic conditions favor protonation, so formaldehyde keeps its H⁺ & doesn’t want to go giving electrons to Ag⁺
The acidic conditions should prevent most Ag⁺ reduction, but but mixing solutions 1 & 2 BEFORE adding solution 3 further shields free Ag⁺ from being reduced
So, at this point, we mainly have a mix of molecules that are kinda minding their own business, BUT, when we add our base, everything will change!
With Part 2, we add the Na₂CO₃ which will “activate” our TSA
Na₂CO₃ deprotonates NH₄⁺ to give us NH₃
NH₃ complexes w/Ag⁺, to form the diamine complex [Ag(NH₃)₂]⁺
Na₂CO₃ deprotonates TSA, which becomes negatively-charged & grabs our complex, shielding it from formaldehyde
Next we’ll add stain to gel & cross our fingers the formaldehyde finds the right targets to reduce (Ag⁺ bound to our proteins)
Then we keep watch… And try to time it so that we stop the staining before it’s too late!
To stop it, we add acetic acid, which protonates things back up and lowers the hydroxide concentrations so the reaction stops (hopefully in time!)
Another cool application of this silver-making magic is Tollens test. You might have run one of these in an undergrad o-chem lab where you’re given an “unknown” substance and then you run tests on it to figure out what it is (my absolute favorite lab in undergrad!). Tollens test helps you distinguish between an aldehyde and a ketone (which has carbon groups on both sides of the carbonyl). Ketones don’t have the power to reduce silver ions, so they will give a “negative” result (no solid silver) but if there’s an aldehyde, you’ll get solid silver plating your tube.
A similar reaction used to be use to test for diabetes, since glucose (blood sugar) is an aldehyde.
For more information:
Chevallet, M., Luche, S. & Rabilloud, T. Silver staining of proteins in polyacrylamide gels. Nat Protoc 1, 1852–1858 (2006). https://doi.org/10.1038/nprot.2006.288 https://www.nature.com/articles/nprot.2006.288
Slideshow by Elaine Randall on SlidePlayer https://slideplayer.com/slide/4366092/ which was super helpful for figuring out what’s in the stains