Some days, I find it really hard to concentrate… my protein that is! One of the perks of starting early in the lab is that you can hog the centrifuge for hours – it’s usually a hot commodity because we do a lot of protein concentration using “centrifugal ultrafiltration” which is just a fancy-dancey way of saying you stick your too-watery protein solution into a membrane-lined tube insert and spin it really fast. The force from the spinning pulls the water (plus salts and other small things) through the membrane, but your protein’s too big to get through the membrane’s pores so it stays put. Sounds pretty boring – and it is – especially when your protein is taking hours to concentrate to the desired concentration… but it’s really important and we do it a lot so today’s a short, practical, post I hope will bore you not…
Protein purification usually involves a technique called column chromatography, where you pass a solution containing your protein of interest (and other proteins) through a series of columns filled with little beads called resin that have different properties and interact differently with different proteins (because different proteins also have different properties), allowing you to separate proteins by things like charge (with ion exchange chromatography) & size (with Size Exclusion Chromatography (SEC) and isolate the protein you want. http://bit.ly/proteincleaning
There are a couple of times during the protein purification process when you want/need to concentrate your protein
- before Size Exclusion Chromatography (SEC) (aka Gel Filtration)
- before freezing your final product
Why before SEC?
A lot of forms of chromatography in effect concentrate your protein for you. Take, for example, affinity chromatography, where the resin is specifically sticky for something special about your protein. Last week we looked at one such form, where I had a His-tag on the end of my protein which binds to nickel bound to the resin. I used this technique again this morning. When I added a “dirty” protein mix, my his-tagged protein stuck, the other stuff flowed through, and then I pushed my protein off (eluted it) with a competitor called imidazole. http://bit.ly/histagpurification
Here, the starting volume does NOT determine the finishing volume – I often start with hundreds of mL of cell lysate (the stuff that spills out of cells when you break them open (lyse them)) and elute my protein with just 20-50ish mL. I can do this because my protein sticks to the column and I get to control when it unsticks and – to a degree – how much volume to get it to unstick with.
But, with SEC, the starting volume DOES matter – because unlike affinity chromatography, with SEC, there’s no sticking. Instead, you inject a sample containing proteins and those proteins get separated by size because they travel differently through tunnels in the beads (bigger proteins are too big to get into the tunnels so they travel a shorter distance and come out before smaller ones). Since there’s no sticking, the proteins that start the journey first (first contact the resin) will have a head start over the proteins that are at the end of your injection, and they’ll end up slightly ahead. How “slightly” depends on the volume you injected. So, in order to prevent unfair advantages (which show up as broad, diffuse, poorly-separated peaks on the chromatograph telling you when proteins elute) you want to keep the volume as low as possible. http://bit.ly/sizeexclusionchromatography
I wanted to take my elution from the Ni-NTA column and run it on a SEC. I know I said you could kinda control your elution volume there, but you have to wash through enough competitor and get all the stragglers, so you end up with more volume than desired. 20mL is WAY too much to put through a SEC column. Instead, I would need to get it down to a couple of mL (note: this is for the big SEC column – when I run smaller columns, I inject much less. the rule of thumb is that, the lower the volume, the better, and don’t exceed ~2% of the total column volume).
So I was gonna have to concentrate. A lot…. So I pulled out (well, down…) the concentrators.
Which one? So many to choose from! Protein concentrators come in many volume-holding-capactities (e.g. 0.5mL, 4mL, 15mL) & molecular weight cut-offs (MWCO) (e.g. 3K, 5K, 10K, 50K). MWCO refers (indirectly) to the size of the membrane’s pores. It’s given in units of Daltons (Da) & tells you molecules below this size can go through (are penetrating) but molecules above this size are retained (are non-penetrating & stay in the top). You want to choose a MWCO smaller than your protein (& anything else you want to keep) but larger than whatever you want to get rid of.
You put your sample in the top chamber & spin it it the centrifuge.
Molecules < MWCO are pulled through the membrane into the lower (waste) chamber, but molecules > MWCO stay in the upper chamber
The bigger the pore size, the faster you’ll reach equilibrium (because if a molecule bumps into the membrane it’s more likely to “bump into” an open space it can get through & doesn’t have to worry as much about “squeezing” through. BUT you want to be careful not to select a size too close to your protein size since the MWCO is an average, so you still might have pores big enough to let your protein through.
Typically, a MWCO “guarantees” that at least 90% of molecules of that size will be retained. BUT proteins have different shapes which MW doesn’t account for (e.g. a long skinny protein might be able to “slither through.” So to avoid losing protein, you typically choose a MWCO 1/2 the size of smallest thing you want to keep. Note: this might remind you of dialysis… http://bit.ly/2KxDEVF
Another important thing to keep in mind is that, since it’s an average pore size and since all the proteins are still able to mix around with one another, it’s NOT useful for separating proteins by size. Ultrafiltration can only be used to separate things that differ by a magnitude of size. So I can separate my protein from salts, but not from another protein.
Also, since we’re on the topic of salts, you can use this as a way to “desalt” a protein and/or switch it into a different buffer – concentrate the protein and then re-dilute it in the buffer you want.
I usually concentrate it in spurts of 15min or so depending on how much concentrating I need to do. In between spurts I use a pipet to mix around the liquid, especially near the membrane, where gunk can build up on the membrane walls and make passage more difficult.
So, after many rounds of this, my protein was finally concentrated enough. And then I ran the SEC. And collected the eluted (now even purer) protein. And…. concentrated it! again!
So, why we am I concentrating my protein again? It’s usually best to store proteins at a fairly high concentration for a couple reasons:
1) I want to give it a more “crowded” solution (like the protein is used to inside a cell). This discourages unfolding (similar to how it’s hard to sprawl out on a crowded bus, but manspreading is easily tolerated when the bus is pretty empty) and helps keep the protein in its proper shape
2) some protein will always stick to the container walls, but w/higher concentration, this has less of a proportional effect on the protein concentration (like losing a drop of water in a pool vs a drop of water from a teaspoon.
But warning! Too high of a concentration can lead to aggregation. So you want to find a “goldilocks” concentration range. This differs for different proteins so, if you’re working with a new protein you might need to test its limits (on small fractions of it so you don’t crash it all out!)
Thankfully, my protein didn’t crash, but my brain is, so sorry for the short post that probably won’t be read by many of you because I bored you halfway through… But hope it was helpful to some people.
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
more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉 http://bit.ly/2OllAB0 ⠀