If you stick a bunch of histidines in a row (about 6 or 8 or so) and over a nickel column flow, that “His tag” will stick so your tagged protein you can “pick,” then compete it off with imidazole – it’s quite a cool trick! IMAC isn’t some new product release from Apple. But it is, at least for biochemists, even more exciting! IMAC stands for Immobilized Metal Affinity Chromatography and it’s a way to help specifically purify “His-tagged” proteins

text adapted from past posts, video new

Histidine is an amino acid – it’s a normal protein letter that can link up to the other amino acids to form long polypeptide chains that fold up into functional proteins. It may be *normal* but it’s also special because it has this property called resonance that I’ll get into which allows it to bind metals like nickel (Ni). If you put a bunch of His in a row, they can team up to really clamp on. So if that Ni is also bound to little beads called resin you can get a His-tagged thing to stick to the resin. We often use this technique to purify proteins which we engineer to have a His tag at the end. ⠀

I will talk more about this later, but basically, since His is a normal letter, we can just stick the genetic instructions for making it onto the end of the genetic instructions for making the protein and the protein will get made with that extra bit without the cell even getting suspicious. Of course, in order to really be useful for protein purification, the binding needs to be reversible. And it is – you can add a competitor called imadazole which is basically just the ring-y part of His to push it off. So let’s look in more detail at what’s going on, as well as some practical tips for using IMAC in the lab.⠀

We’ll start with what makes His sticky…⠀

Atoms link up by sharing pairs of electrons – you need 2 for a single bond & 4 for one of the shorter, stronger, double bonds. But what if you don’t quite have enough? You might want to join an electron commune! (otherwise know as resonance/electron delocalization/conjugation). Histidine’s a fan of this. Histidine is aromatic. That doesn’t mean it smells nice, it just means that it has a ring where, after they’ve “spent” 1 electron each on the bonds to their neighbors the atoms in the ring donate their “extra” into a communal shared stock. Those atoms that opt into this commune get to share, and this leads to electron delocalization above and below aromatic rings, kinda like a donut. ⠀

His’ side chain has a 5-member ring and 2 of this ring’s corners (yeah, it’s more of a polygon than a actual ring…) are nitrogens (N) instead of the “usual” carbon (so we call it heterocyclic – hetero meaning different). These Ns are able to join the commune because they have a loan pair of e⁻ that can take “role” of double bonds in resonance structures in providing “extra” e⁻ ⠀

Now let’s get back to that aromaticity – His’s aromaticity attracts ➕ charged metal ions. And this is really useful because metals are really useful – both in your body and in the lab. A lot of enzymes (molecules (often proteins) that facilitate chemical reactions) bind metals & use them as “helpers.” Why are metals helpful? For one thing, they’re really big & multiple parts of the enzyme can latch onto them so it acts as a sort of coordinating center to keep the enzyme’s structure together. ⠀

Metals are so big because they have huge electron clouds – the places where electrons whizz around their atomic nuclei containing the positive protons reigning them in. When metals bond to other metals in metallic bonds, they kinda just merge some of their clouds into a vast, communal electron sea. And when electrons move throughout the sea you have electric current -> this is why metals are good electrical conductors.⠀

But when metals bond to nonmetals, they often do so as coordinate covalent bonds – these bonds form when 1 atom shares 2 electrons with a metal ion in the middle (in normal covalent bonds each atom shares one) – we call the result a complex and you can learn more about them here: https://bit.ly/2BDBml0

In order to complex with a metal, an atom has to have a lone pair of electrons they can share – like nitrogen!!!!!. If a molecule has more than one atom with a pair to spare, they can “bite down” on the metal in multiple places -> polydentate. We call such multiple-toothed-biters chelators.⠀

His’ Ns can act as electron pair donors in coordinate covalent bonds, so His is able to take a bite. And this can hold metals in place in the active site of enzymes where those metals can give & take electrons to help promote reactions & stabilize awkward reaction intermediates. ⠀

If you have a lot of His in a row, it’s like having a polydendate (many-biting) thing – one of those chelators. So if something has a string of His it can bind to metal tightly. And we can take advantage of this to help us purify proteins. ⠀

If we want to study a protein, we usually need a lot of it. And we need it pure. So we express it recombinantly – this means we take the gene with instructions for the protein we want, recombine it with a piece of DNA that makes it easier to work with called a vector, then stick that vector into cells (often bacteria or insect cells) to make the protein for us. They’ll make it, but it’s up to us to get it out of the cells and purified.⠀

So we want a way to make it easier to isolate just the protein we want and not any of the other stuff. A way we usually do this is through column chromatography. I flow a solution (mobile phase) containing my protein & contaminants I want to get rid of through a resin (lots of tiny beads) that’s held put in a column (solid or stationary phase). This resin interacts w/different things differently. You want it interact w/your protein one way & the proteins you don’t want another way so that they get separated. Much more on this here: http://bit.ly/bbproteinchromatography

We can use different types of resin to separate based on different properties – properties that are “inherently normal” to the protein – like size or charge or something we’ve added on through recombinant expression trickery. Affinity chromatography uses resin that recognizes something really specific – usually an affinity tag we’ve added onto the end of the protein (by putting the genetic instructions for it before or after the instructions for our gene in the plasmid). Because affinity chromatography is recognizing something “unnatural” and highly specific, it can (hopefully) remove most contaminating proteins – but not all.⠀

A common affinity tag is a His tag, which is just 6 or 8 Histidines, which will bind to a Nickel (Ni) or Cobalt (Co) coated column (Immobilized Metal Affinity Chromatography; IMAC). Other proteins have Histidines too. But not that many in a row, so your protein will bind preferentially. It’ll hog the column and the other proteins will flow through.⠀

Then you need something that will outcompete the His tag to get your protein off. Bring on the imidazole. It looks like His, so you can flood the column with imidazole to push the His-tagged proteins off. But before you flood it, you wash it with low levels of imidazole to remove non-specific binders that are just binding cuz they happen to have a lot of Hises.⠀

So you go from ⠀


resin:Ni:His-tagged protein⠀

then resin:Ni:imidazole + His-tagged protein⠀

In the beginning, when you’re loading your column, flowing through all that gunk, your protein is able to push off the imidazole because Imidazole mimics a single His so can only take a single bite whereas your tagged protein has a lot of them. When you are ready to push your protein off (once you’ve washed off all the other stuff) you can do it with imidazole, but you have to add a ton of it since your protein’s a better binder – you have to go with the “flood it out” strategy.⠀

Nickel isn’t the only metal you can use. Another one I’ve tried is Cobalt. The most obvious difference with cobalt is its color – it’s pinkish red. The less obvious difference has to do with its “pickiness” (specificity) (cobalt has a higher specificity) and “stickiness” (affinity) (cobalt has a lower affinity), which have to do with its subatomic makeup. Co & Ni are both “transition metals” – you find these in the center of the periodic table and they’re good at giving and taking electrons (can have multiple oxidation states – more here: http://bit.ly/31MPb9w )⠀

They’re really similar in a lot of ways, and both bind the beads and your protein. But Ni has 1 more proton (giving it an atomic # of 28 to Co’s 27). And they have slightly different “tastes” when it comes to molecule dating. It’s kinda like every time a protein meets the metal the metal has the choice of “swiping left or right.” The chances depend on how attractive the metal finds the protein. The higher the affinity, the more attractive. But if it finds everyone attractive, you have low specificity.⠀

If your protein’s less attractive, you’ll need more metal-coated beads so you have more of those encounters – even though each encounter still has a lower probability of sticking, if you have a lot of them your chances improve. And as long as its affinity’s a lot better than the contaminating proteins – even if it’s still not great – you’ll get the binding to be almost all your stuff. Nickel finds His more attractive than cobalt does. So, cobalt has a lower affinity but higher specificity because it’s “pickier” and its pickier because it has stricter spatial requirements – the Hises have to be next-door neighbors like you have in a His tag or at least specially spatially positioned. More about cobalt here: http://bit.ly/2kuAAic

Some practical notes…⠀

The resin we use for IMAC is usually beads made of agarose coated with chelators – the one I’m using is NTA – nitrilotriacetic acid (NTA), which is a tetradentate chelator (4 sites). You might have seen cobalt resin advertised as TALON – that’s just a brand name for a slightly different linker – a patented carboxymethyl aspartate. Nickel and Cobalt offer 6 places to bite, so you still have 2 spots left for your protein to bite. ⠀

So that His-metal bite is weaker than the resin-metal bite (2 bites vs. 4), so you can get your protein to come off (elute) without stripping the metal off, just by adding a His mimic called imidazole that acts as a competitor. Imidazole mimics a single His so can only take a single bite – it’s not a chelator so you have to add a lot of it. ⠀

If you add a stronger competitor like EDTA, which *is* a chelator and can bite down on all 6 of the metal’s bite spots. you can get the metal to come off too. So you can turn your blue resin un-blue. So you can strip the metal off and give it a wardrobe change into a different metal – like cobalt. (Or you can just “recharge” it with the original metal)⠀

Periodic charging (adding more metal, typically by flowing through a metal-chloride solution) is needed because of “leaching.” Leaching is when the metal comes off the beads, not just your protein, and when you don’t want it to – this is more of a wardrobe malfunction! Nickel is more likely to unintentionally flash someone because it can actually bind the resin in a couple different ways – it can form 2 different coordination complexes.  One of these complexes forms a nice little pocket that’s ideal for tag-binding with Ni bound to 3 carboxyl groups & 1 N. But the other complex has a flat (planar) structure, where nickel’s bound to 2 carboxyl groups & 1 N. This doesn’t bind the Ni very tightly, so the Ni can leach off.⠀

With cobalt, the linker bites the Co harder, but the protein bites it more weakly. So the Co is less likely to leave the column, but your protein is more likely to leave the Co. So – Ni & Co both will bind the beads and your protein, but Co binds the beads tighter – so it’s less likely to leach off the beads (this is what I was aiming for). Co may bind beads tighter but proteins, not so much. It has a lower affinity than Ni for His, so, unless you use more resin, it might yield a lower yield. But, since it binds non-specific stuff worse too and that stuff bind worse to begin with, you end up with a purer protein product!⠀

The basic protocol I use for recharging resin is:

  • strip w/0.25M EDTA, 0.5M NaCl
  • wash with water
  • regenerate with NiSO₄
  • wash with high imidazole buffer
  • wash with water

fun sidenote: When I was looking online to see if it’s a-okay to strip & swap the metal from the Ni-NTA resin we buy, I saw some references to cobalt being blue. And isn’t “cobalt blue” even like a crayon color? But the cobalt I found on our lab shelf was cranberry-colored. That’s because cobalt’s color depends on who it hangs out with. 

I prepared the cobalt solution from Cobalt Chloride hexahydrate CoCl₂·6H₂O- the “cobalt chloride (CoCl2₂) part” tells me that, in this salt form, each cobalt atom hangs out with 2 chlorine atoms (the Co has a +2 charge that’s neutralized by the -1 charge of 2 Cl’s). And the “hexahydrate part” tells me that each of these trios is surrounded by 6 water molecules.

And apparently when cobalt chloride is “wet” it’s pink or purple but when it’s “dry” (anhydrous) it’s blue. When you get the “anhydrous” form wet it first forms a dihydrate CoCl₂·2H₂O which is purple. And when you add even more water it turns pink. So cobalt can actually be used to indicate the presence of water and they make like water test strips out of it to test pipes for leakage, use it to indicate desiccator capacity, etc. 

Note: Another affinity tag I use is a Strep tag which binds streptactin resin. I use the Strep tag when I do insect cell expression, but the His-tag with the bacterially-expressed stuff. His is underrepresented among the protein letters (amino acids) – there are 20 (common) ones yet His only makes up ~2% of amino acids you actually find in proteins. And, you’re more likely to find adjacent ones by chance in eukaryotes than prokaryotes – less contaminating proteins binding non-specifically when used for bacterially-expressed proteins⠀more here: https://bit.ly/streptagaffinity

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