Using DTT? How could you tell? I recognized that distinct rotten-egg smell! Can the bumbling biochemist reduce REDUCING AGENTS into a post? Maybe if she focuses on the ones biochemists use most – DTT, TCEP, & BME keep the REDOX environment in our artificial buffer sea similar to that intracellular proteins see. To understand why let’s go for a dig into the redox world of OIL RIG!
OIL RIG is a mnemonic we use to remember Oxidation Is Loss (of electrons) and Reduction Is Gain (of electrons). Electrons are negatively charged but positively cool! They’re little subatomic particles that whizz around the dense, positively-charged, nucleus of atoms.
Different atoms have different numbers of electrons. They keep most of them to themselves, held in by the positively-charged protons in the nucleus, but the electrons furthest away from the nucleus (valence electrons) feel the pull less, so they can play the field more to get to their “ideal” For some atoms, losing electrons gets them closer to ideal and for others, gaining electrons gets them closer. So, in “rig-ged reactions” the molecules conspire so each gets what they want.
Atoms can lose electrons – which we call oxidation – the OIL in OIL RIG. And when they do, the electrons have to go somewhere – and the somewhere they go to is another atom. So that other atom gains electrons, and we call that reduction – the RIG. Together, we call these redox reactions and you can’t have 1 without the other.
One molecule, the reducing agent, or REDUCTANT, “wants” to get rid of electron(s) and the other molecule, the oxidizing agent, or OXIDANT “wants” to take them.
This is easiest to see in reactions with metals changing charge, but it’s often less obvious in reactions taking place between biochemical molecules. The important thing to remember is that instead of losing actual electrons, you can think of oxidation as losing some electron density – electrons don’t stay in place and when we draw them in pictures, we’re just showing where they’re most likely to be. So even if an atom looks like it still has the same number of electrons, if it goes from keeping them to themselves to sharing them with something else it’s “losing” some of the time the electron’s hanging out with it.
Often in biochemistry, the oxidant is a sulfur (S) or oxygen (O) which often have unshared “lone pairs” of electrons they’re willing to share. Even though they don’t really “lose” the electrons they’re sharing, since they’re sharing it’s like they’re losing 1.
But once the oxidant takes an electron it becomes reduced and once the reductant gives the oxidant the electron, the reductant becomes oxidized and reluctant to act as an oxidant! So it’s not like something’s always in the “I want to reduce” state or alway in the “I want to oxidize” state. Their “mood” depends on their “redox environment” – the amounts of reductants and oxidants around them.
Our cells maintain a reducing environment in part to protect against oxidative damage. Oxidants like reactive oxygen species (ROS) can break up existing molecular relationships, doing things like breaking DNA if they’re not kept in check.
Cells control the redox environment by stocking up on reducing agents like glutathione (you might remember this from my post on GST-tagged proteins). There, we use glutathione to compete for binding to a GST tag, not for its reducing properties. Glutathione (made from 3 protein letters (amino acids) – Glu, Cys, & Gly) acts as a “Redox buffer” 👉goes back and forth between reduced (GSH) & oxidized forms(GSSG) 👉 2 glutathione linked by disulfide bond.
Proteins (molecular machines) are made up up of chains of building block “letters” called amino acids that are like charm bracelets. Amino acids have a generic backbone (chain link) 🔗 that allows any amino acid to connect to any other amino acid as well as a unique side chain “charm” that sticks out 🌟 Charms have different chemical properties that allow them to interact in different ways w/one another (important for the protein to fold properly) & w/other molecules (important for intracellular interactions) 👍
The reducing power of glutathione comes from the Cys, whose “charm” is a -CH₂-SH group 👉 -SH is called a THIOL (it’s alcohol’s (-OH) sulfur CYSter) (more on alcohols here: http://bit.ly/2QWKWXp). 2 Cys (either within the same protein or between 2 proteins) can link together (protein)-SH + HS-(protein) 👉 (protein)-S-S-(protein) to give you a disulfide bond (aka a disulfide bridge or cross-link) & now you call the Cys’s cystIne 👍
Unlike other charm-charm interactions, which are just *attractions* based on charge (or partial charge) differences 😍, this is a covalent bond, so it’s strong 💪🏻 (and good for sturdying-up secreted proteins that have to live outside the comfort of the cell), BUT it’s not quite as strong as the covalent bonds in the protein’s backbone (linking the chain links 🔗) so you can split them back up without splitting up the chain links 👍
When cysteine’s link up as cystines, they share their “extra” e⁻ so they’re “losing” e⁻ 👉 being OXIDIZED (lose the “e” in the name (and also “e⁻”)) 👍
When they split up, they get the e⁻ they’d been sharing “back” 👉 they’re REDUCED 👍
DISULFIDE BONDS or “cross-links” can form between 2 Cys residues in the same or different proteins, & can be important for maintaining some proteins’ shape – but only if the right ones form. And because these bonds are stronger than the normal intermolecular bonds, they’re harder to break up, so it’s important you get it right.
If a cysteine residue gets reduced, it gains electrons – and the negative charge they bring – so it becomes nucleophilic – seeks out the positive charge coming from a neighboring atom’s nucleus. So you want to make sure the neighbor it finds is the neighbor it’s supposed to bind!
If you have an oxidizing environment, proteins can panic like it’s the end of the world & hook up with the nearest available thing instead of waiting for their “soulmate.” This is actually how perms work – you use reducing agents to break up the disulfide bonds between strands of keratin in hair – then you add oxidizing agents to reform them, but you do this when the hair is physically curled so that the nearest neighbors are the ones that will make the hair stay curled
When we purify proteins, we usually try to keep them in an environment that’s close to what they’re naturally found in. Evolution’s had years and years of natural selection to adapt these proteins to be happiest and most active in that environment.
When we’re studying proteins that are normally inside of the general “inside” of cells, the environment we’re trying to mimic is that of the cytoplasm. There’s a lot of stuff going on in there, and we don’t want to add too much complexity to our buffers, especially since the actual contents of the cytoplasm varies from cell type to cell type and even cell to cell.
So, in our mimicking we try to stick to the minimal amount of things that will make our protein happy, soluble, and active. The artificial “seas” we put our proteins in in the lab are called BUFFERS. The name refers to the pH-stabilizing component we put in there.
pH is a measure of how acidic (proton-rich) or basic (proton-poor) a liquid is. Neutral’s 7 – lower’s more acidic & higher’s more basic. Buffers are molecules that can give and take protons to keep pH constant. Some ones we often use are Tris, HEPES, and sodium phosphate, and you can learn more about them here: http://bit.ly/2IT67pC
We usually buffer the pH around 7.4-8, which reflects cytoplasmic conditions.
Another thing we need is salt. A couple common salts we use are NaCl (sodium chloride, aka table salt), KCl (potassium chloride)
We also usually add a reducing agent. Instead of using glutathione, which oxidizes too easily, a few common reducing agents we use in the lab are DTT (DiThioThreiotol), TCEP ((Tris(2-Carboxyethyl) phosphine hydrochloride)), & BME (β-mercaptoethanol, aka 2-mercaptoenthanol). They all can reduce disulfide bonds, but they also have some different properties that make them better or worse in different situations.
SMELL: DTT & BME both have that rotten-eggy smell because they both have sulfur in them (the “thio” and “mercapto” give this away). And if you don’t like the smell of DTT (which I don’t think is that bad though maybe I’m just use to it) you don’t want to smell BME – it’s even worse! But TCEPT doesn’t have sulfur so your nose is safe.
COMPATIBILITY: EDTA turns Ni-NTA resin white and NiSO4 turns it blue – but if your IMAC resin turns brown, you likely have DTT hanging around
DTT can turn Ni-coated resin, like the Ni-NTA we use for IMAC, brown – this is because it can reduce Ni. Unlike EDTA which actually removes the Ni from the column, this “browning” usually doesn’t actually interfere with protein binding because it’s interacting with the Ni that’s “uncoordinated” – only loosely bound. But it can – and it’s not pretty – but is pretty disconcerting…
A way you can use DTT (at low levels) with this resin without it turning brown is to, before you let the column see DTT, pre-wash it with a buffer with a high concentration of imidazole. The imidazole will wash that loose Ni from the resin. (and you don’t really want that loose Ni there anyway right?)
TCEP absorbs less light at 280nm (one of the wavelengths we use to measure protein concentration)
size: size-wise TCEP > DTT > BME. You can tell by looking at their molecular weights (M.W.) which tells you how much 1 mole (6.02e23 molecules weighs (in g). TCEP has a M.W. of 250.15 (250.15 in 1 mole), DTT’s 154.25, & BME’s 78.13. Why care?
Because TCEPs bigger it has a harder time getting to disulfide bonds that are present “inside” a protein – so it can readily break up intERmolecular bonds (between different proteins) and thus break up dimers but it leaves intRAmolecular bonds (within a single protein) so it doesn’t make them unfold. But, as long as you don’t crank the amount of reducing agent too high (as long as you’re keeping reducing conditions to those like the ones in the cell) you should be ok on this front regardless – if your protein’s that sensitive it wouldn’t be able to stay folded in the cell.
Speaking of which, proteins that do have vulnerable disulfide bonds that need to be kept go through alternative processing pathways where they’re made & shuttled out of the cell (into the oxidizing extracellular environment) in protective membrane-bound vesicles.
COST: TCEP is much more expensive. It might not seem like a big deal price-wise if you don’t need much, but when you have to do things like dialysis where you need liters and liters (I had to dialyze against 8L the other day) that price difference really adds up
STABILITY: DTT’s not very stable, so we usually add it “last minute.” What I normally do is, for buffers I make a lot, I make a 10X stock. Then when I want to use it, I just dilute it 1:!0 to get to the working concentration (1X) and add DTT then. TCEP isn’t very stable in phosphate buffers – esp. at neutral pH
more about redox: http://bit.ly/2P4FekB