At the end of a protein purification, you often are really exhausted and want to take a nap – but first you have to prepare your protein samples for *their* nap! You need to get them all snuggled up with cryoprotectants like glycerol, and then knock them out with flash-freezing in liquid nitrogen before the ice monsters can attack! (Water ice crystals form and harm your protein). That’s right – you need to vitrify your samples! The concepts are really similar to what you might do to prevent ice from forming in pipes, so I thought I’d pipe in with a “supercool” video post based on a prior post. 

So, video new, text adapted from that longer past post which you can find here: http://bit.ly/flashfreezingdance



If you you imagine a molecular “prom dance” – in a liquid the dancers are free to glide around & swap dance partners – they just can’t leave the dance hall. But in a solid the molecules can only stand in place and do that awkward sway thing with their partner(s). The difference between them is that in a solid the molecules don’t have the energy to “un-hug” their dance partners and go find someone else. What the molecules really want to do is leave the dance hall all together (escape as a gas) but that requires making their way to the exit without having new dance partners ask for a dance. So that takes a lot of energy.

The stronger the partners are hugging the more energy is needed to un-hug and water’s really huggy because the oxygen doesn’t share fairly – it hogs electrons, making it partly positive & the H’s partly negative – and opposites attract so they like to hang with oxygens from other water molecules.

Molecules would rather be on their own (where they have more entropy (randomness/disorder – basically they can move more ways and this makes them happier), but if they have to be there, and they have to get stuck in the super slow dance of solid-ness they’d rather be stuck with dance partners they like. So if they have “advance warning” that a slow song is coming they can link up to their preferred partners and doing so generates a rigid orderly arrangement of molecules we call a crystal.

And that linking-upness may require pushing out the “noncool kids.” So with slow freezing you end up freezing with different “cliques” in different areas of the dance hall – a “heterogeneous” mix instead of the nice evenly-distributed molecular crowd that we call a “homogenous solution”. And if you’re dealing with water, the molecules literally get “pushed” because water expands when it freezes (even if the water’s inside pockets & channels of your protein!)

But if you can switch on the slow song without that advance warning, the molecules get stuck in place – this is the principle behind “flash freezing” And if you add cryoprotectants like glycerol that act like “dance chaperones” that break up the dance partners and make it harder for partners to find each other, you buy yourself more time

In a solid, molecules are “stuck in place” & only have enough energy to vibrate (as opposed to liquids where they can slide around & gases where they can go wherever they want). In some solids (like ice & diamonds), the molecules are arranged in an orderly lattice, & we call this a crystal or crystalline solid

But going doing that whole liquid -> crystal thing (crystallization) takes a lot of coordination – It requires nucleation. Kinda like an unplanned flash mob dance – you need “trendsetting” molecules to freeze together in the “right” orientation (nucleate) to form a “seed” & then others join in & those others have to be able to find that seed & get there before they run out of energy themselves.

So if you don’t want a crystal to form you can:

– make them too tired to find each other – molecular movement requires energy – and heat is a form of energy – so if you lower the temperature, you sap their energy so they’re “stuck with swaying in place”

– not give them enough time to find each other (cool it really quickly)

– make it harder to get there – use viscous (syrupy) liquids that are hard to move through

– make it harder to find each other – add cryoprotectants like glycerol that act as “dance chaperones,” breaking the molecules up and making it harder to “find Waldo” – the “costliest” part of crystallization is that initial trend-setting (nucleation) – so that’s what you really really want to make it hard to do

– make them like each other less –  how much molecules like each other versus the liquid their in or other molecules around depends on things like the pH and salt concentration of the liquid – we often take advantage of this – but in the opposite way – in protein crystallography. There we where we *want* our *proteins* (but not the water in them) to form nice orderly crystals so that we can shine x-ray beams at them and have those beams bounce off a uniformly-arranged object so we can work backwards from them and figure out what the protein looks like. So we can gradually change pH and salt concentration and stuff to get the molecules to like each other and/or hate the water more.more here: https://bit.ly/gettingcrystals

But for normal protein work, I don’t want the proteins linking together either. Instead I want to “freeze” them in place – basically just take away their energy – I want them to be as much like how they are now when I wake them up as possible. So what I want is a disordered, amorphous solid.  We call this a glass & the process of forming it is vitrification 

The glass transition temperature (Tg) is the range of temperatures in which a glass can form & it’s lower than the freezing temperature (when you go liquid -> crystal). So in order to get to it you have to quickly skip past the freezing point to get to a supercooled state (which you can do because of the coordination required for crystallization).  Cryoprotectants help because they lower the freezing point & raise the Tg (so you have less time in “danger zone” between them) – this is because now, if it wants to form a crystal the water molecules must “push away” the glycerol molecules in between them before they can link together in their exclusive “clique.”

We saw cryoprotectants at work a while back when we looked at why salt is used to prevent ice forming on roads. Salt is good at that because freezing point depression is a “colligative property” meaning it depends “only” on the number of dissolved particles, not their identity. And table salt (NaCl) is a sort of 2-for-one. It doesn’t just dissolve in water (get a full water coat), it also dissociates, meaning that the Na⁺ and Cl⁻ ions come apart, giving you twice as many molecules as you put in, and thus twice the freezing point lowering. http://bit.ly/freezingpointdepression 

But we don’t want to oversalt our proteins when we freeze them because that could mess up the protein interactions. Thankfully there are alternatives. You want to add something that’s more “water-like” in terms of not disrupting the system (but less water-like in terms of “stickiness”). Basically, you want something kinda “blah” – and small so we can pack lots of it in there (colligative properties, remember). Small polyols (chemicals with multiple -OH groups) like glycerol fit the bill and thus are commonly used as cryoprotectants.

Glycerol is what I use when I’ve finished purifying a protein with a series of chromatography steps (much more in prior posts like http://bit.ly/bbproteinchromatography). After the final step (typically size exclusion chromatography (SEC), which separates proteins by size) I concentrate my protein using one of those spin ultrafiltration thingies I talked about before. I use glycerol as my cryoprotectant because it’s water-like so it’s not a big shock to my protein – but it’s less sticky than water. Some proteins, especially some enzymes, need high glycerol concentrations, but I use 10% for most of my stuff. Glycerol is super viscous (syrupy) – this helps it cushion proteins, but makes it really hard to pipet. So instead I pipet from a 50% or 80% glycerol stock.

And then I “flash freeze” them – and remember, when I say I freeze my proteins I’m not *really* freezing them – my proteins are way too cool to do that to – instead I “supercool” them before I store them away for safe keeping in the -80°C freezer by dunking tubes of protein in liquid nitrogen to race through the freezing point & outrun ice formation. 

Liquid nitrogen is sometimes abbreviated LN2 , referring to the fact that the atoms of nitrogen (N) are present in pairs (it’s diatomic). Nitrogen has a freezing point of −210 °C (−346 °F) so it can exist in a liquid form at very very cold temperatures (temperatures at which water w/a freezing point of 0°C (32 °F) would be frozen solid). So even though I’m using it as a liquid, it’s really really cold! (remember to take safety precautions – freeze your samples not your skin!)

With water, I’m worried about freezing, but with LN2, the bigger concern is *boiling*! When it comes into contact with warmer things (such as my samples), the heat from the warmer thing is transferred to the LN2. As a result, the sample is rapidly frozen, but the LN2 evaporates into N2 gas. This looks really cool but also means you’ll need more of the liquid form than you expect (our lab goes through a ton of it!). more here: http://bit.ly/30JJBni

A great thing about vitrification is that, since the molecules stay where they were as a liquid, you don’t get the expansion you get with ice & the solutes can stay put too so the environment stays the same. So, (hopefully) you can “wake up” the protein and it’ll act the same. Well, it’s never really quite the same – when you thaw a frozen sample, you’re adding energy that lets the molecules break free from one another & seek out new binding partners. But if you remove the energy again, they get stuck in place again (re-freeze). You can keep doing this freeze-thawing, but each time you do this you have more chances of crystals forming and damaging your sample. So you want to avoid multiple freeze-thaws by making aliquots, which are like “single-portion” packages so that when I want to use some I only have to unfreeze what I want to use.

For more practical protein-purification posts (and background/theory), check out the new page on my blog where I’ve collected some of my protein purification posts. http://bit.ly/proteinpurificationtech


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