Now how did this happen...
April 10, 2011 3:21 PM   Subscribe

ProteinFilter: Anyone ever worked with a protein that forms a filmy mess after purification?

I'm fairly sure that this is a bacterial contamination issue. However, the protein shares loose n-terminal homology with spectrins and has been shown to be membrane associated. After concentrating down and spinning out the gunk, the yield of protein is roughly halved, so I'm a little suspicious as to whether or not it actually bacteria I'm seeing.

Purified from BL21s, denatured in urea, purified in urea, renatured in Tris/PMSF/Azide/BMe buffer then concentrated on a filter column.

Just wondering if anyone else has had a similar experience in the lab at any point and if it turned out to be bacterial in nature (probably 99.9% of cases) or some really strange protein behavior.
posted by Slackermagee to Science & Nature (13 answers total) 1 user marked this as a favorite
That sounds a lot more like the protein to me. Maybe you could say a bit more what you mean by filmy mess? Have you tried resuspending the filmy mess/pellet in SDS loading buffer to see what is there?

I have a protein which, when concentrated, aggregates in the form or weird stringy things, and I'm sure it's the protein.
posted by lab.beetle at 3:59 PM on April 10, 2011

Yeah, I agree with lab.beetle. After the concentration step, is it still in BME? If not (and it has free cysteines) it could be oxidizing into the big mess. I've definitely seen that happen before. How long after purification does this mess appear?
posted by Durin's Bane at 4:02 PM on April 10, 2011

Response by poster: I concentrate about 20ml with 200ul of BMe added straight in for good measure and its still giving me this stringy, filmy mess. I will dissolve some for a gel I'm running later on. Its less gooey and more... I dunno how to describe it, sheet like? Its a sheet of colloid I guess would be the best description.

Oh, and it may have 1 or 3 free cysteines.

As for how long it takes, I dialyze in renaturing buffer for 24 hours, so sometime after purification and 24 hours later (and there's a good amount of BMe in that buffer too...)
posted by Slackermagee at 4:12 PM on April 10, 2011

Your problem is protein aggregation. Are you doing a denaturing prep because the protein goes into an inclusion bodies?

Second since you say it may be membrane associated, try adding a low percentage of surfactant such as triton 100 or NP-40 (0.05 to 0.1% NP40 is your friend)?

The next thing is to take your protein and try moving the pH up or down. Also changing the salt concentration can do wonders for solubility - start with a nice low KCl concentration and go up slowly.

Is this your protocol or are you following a published protocol?
Is this a GST (or other) fusion? 6xHis purification? That sort of thing?

If you're using spin concentrators, try to avoid using the ones where you spin down through the membrane.
posted by sciencegeek at 4:24 PM on April 10, 2011

Response by poster: 6xHis is the tag (n-terminal) and it definitely goes straight to inclusion bodies in bacteria. I've shown it to associate with lipids in the past and developed a set of storage conditions to keep it nice and happy (monodispered and monomeric) in solution with 2cmc of non-ionic detergent.

This is a new purification procedure for me now, usually I renature then purify, but to avoid what my prof suspects is enfolded contamination I'm doing the whole thing in urea.

When purifying after renaturation I've never had this aggregation/film/goo problem (I'm hesitant to call it aggregation as its just too... loose? I've had it aggregate and crash on me before, turns all to flakes and snow and settles out. This seems very different).

I've not played with the pH, probably something to try with the DLS downstairs.

I'd love to concentrate and renature in the better surfacants that are available but with a new set of lipid pulldowns as the next objective, this might interfere with what I'm planning.

Salt is, unfortunately, definitely not the best thing for this protein. DLS results were fairly conclusive about that, any amount of salt spreads out that nice, lovely monodispersed peak.
posted by Slackermagee at 4:59 PM on April 10, 2011

I know it would be a pain in the butt to change things now, but we've had success in using TRX/GST fusions with a TEV cut site. Then you can do the purification without as much fear of things crashing out. TEV is nicely specific (unlike some of those other more promiscuous proteases) and with a HIS tag on the fusion protein the purification of your guy from the TRX/GST is pretty nice and quick to boot.

Alternately, do the renaturing with detergents and then run it over biobeads to get rid of the detergents. I've not done lipid pulldowns so I don't know if this is a stringent enough way of removing the detergents.

Then again, whatever enfolded stuff you're getting with your old protocol might be necessary to keep this protein happy: ie those lipids? Mass spec might be your friend here - of course this all depends on your access to good mass spec.

Personally, I'd go for a fusion protein - you can start cloning now while you play with other variables.

Also: what does a silver stain/colloidal look like? Do you see significant other protein stuff in your sludge? Does your sludge respond to adding lipids by any chance?
posted by sciencegeek at 6:07 PM on April 10, 2011

Seconding the silver stained gel idea. Load heavy - say 10-20 µg. Maybe do a western blot if you have (or can get) a polyclonal against your target molecule. A good purification (OK, what I think of as a good purification, but we inject things into people) will have one band, maybe one or two fragment or truncation bands. If your gel looks like ladder inspection day at the fire-house you have something living in there.

You can get antibodies against E. coli lysate from a couple vendors if you really want to be sure.

From what you describe (I'm picturing a sort of gloppy viscous mess) my gut reaction is that it's not as refolded as you'd like to believe.
posted by Kid Charlemagne at 7:21 PM on April 10, 2011

Left field, but field-work validated idea.

Try adding some DNAse (try Invitrogen) to your lysis buffer (assuming it's compatible with the DNAse - but DNAse *oughtn't* to be compatible in the first place). I use 2ul of their standard stuff in each mL of lysis buffer when using an SDS containing protein lysis buffer. The same goes for dissociating (papain, mostly) cortical tissue for culture.
posted by porpoise at 8:48 PM on April 10, 2011

OK, couple of questions.

I'm confused why you think you have bacterial contamination - if you've done a urea denaturing purification and have the protein stored in an azide solution, what exactly that is alive is going to be in that solution?

What is the species your protein sequence is found in? If you're for example, purifying an E. coli protein using E. coli as the expression host, you're going to for sure purify a heterogeneous mixture of protein complexes etc., and if your protein serves a function similar to spectrin then probably some membrane associated bits will come down with it also. However, again, the denaturing purification should be more than enough to get rid of co-purifying junk.

Do you need to have the n-terminal region included in your expressed sequence? A very good idea when having problems with protein solubility is to identify chunks that are responsible for your woes and deleting them.
posted by StrangerInAStrainedLand at 10:06 PM on April 10, 2011

Response by poster: Science Geek: Actually, that's what were doing :))) Love TEV, so much so that we've got a a freezer box of the stuff right now.

I may silver stain depending on how the CB for the current prep comes out. Going for a more stringent wash on the column, hopefully the fainter bands (current level of purification is hovering around the 95% mark) will have washed away. If it looks good by CB I'll throw a new gel in the silver stain.

Kid Charlemagne: It doesn't gloop when I renature a lysate and not purified protein though... I'd expect it to be the other way around in this case. More updates though which ought to help: After getting it into the nice 2cmc buffer it's still slowly crashing out over time. Gotta check on a BCA tomorrow but in these two preps I may have hit the solubility limit (lots of turbidity... goddamn).

Definitely not bunches and bunches of strong contaminant bands though, regardless. Its all a slight haze in the background with optimized contrast and ~20ug in the well.

Porpoise: Don't think its a DNA issue, but its worth checking out, thanks for the tip!

StrangerInAStrainedLand: My first reaction to the goop was bacteria, and you're spot on with the buffers. I rather like to doubt myself though. Oh, and its human.

In general this is going onto crystal plates as soon as it hits a level of purity my prof approves of. I've tried MBP fusions and GST fusions in the past and while they work, the protein disappears mighty fast after TEV cleave of the tag (and the cleavage itself with larger tags is just terrible...).

I'd love to chop off the c-term, but like I said, this is for a crystal trial and my prof is in love with the idea of getting a picture of the whole thing. Maybe if I can get an NMR showing c-terminal disorderedness....

Anyways, Thanks for all the recommendations so far!
posted by Slackermagee at 10:19 PM on April 10, 2011

In general this is going onto crystal plates as soon as it hits a level of purity my prof approves of. I've tried MBP fusions and GST fusions in the past and while they work, the protein disappears mighty fast after TEV cleave of the tag (and the cleavage itself with larger tags is just terrible...).

I'd love to chop off the c-term, but like I said, this is for a crystal trial and my prof is in love with the idea of getting a picture of the whole thing. Maybe if I can get an NMR showing c-terminal disorderedness....

Some science advice and some grad school advice:

When trying to crystalize a protein, do as the high throughput structural genomics guys do. This means making several constructs with variable length N and C termini based on disorder/order predictions, sequence conservation etc. Also, many eukaryotic proteins are like several globular domains attached by linkers, and its very very difficult most of the time to get that kind of thing to crystalize. Call me pessimistic but you'll have a much greater chance of success when going for a single domain. Which brings me to...

You'll have more success in grad school if you can get out from under your PI's thumb a bit. Mine can be a bit overbearing as well, and like most trends toward "control freak" behavior. The best way I've found to get around this is to simply take the initiative and do many things that I think may or may not work. Some of which actually do work, and those he hears about. PIs are very smart people generally, but grad students are also. Go forth, and make constructs!
posted by StrangerInAStrainedLand at 10:36 PM on April 10, 2011 [1 favorite]

Do you want to give us an accession number so we can take a look at this protein? No problem if you don't want to for reasons of privacy/competition/etc. I'd also like to see putative domain structure if you've played with it enough to have that - otherwise, I can just run it through SMART or something.

About a year ago, I spent a bunch of time working with a recalcitrant protein which didn't like being soluble. One of the things I figured out was that it resented being frozen - even flash freeze in liquid nitrogen - so I had to purify it quickly and use it quickly. It also hated being concentrated - ended up using a vacuum concentrator because I lost more than 50% using spin columns.

Things I did learn that might be helpful (some of these might be obvious): TEV can cleave under a wide variety of salt and pH conditions and this can help you keep your protein happy for more of the purification. If you've got a cofactor of any sorts keep it in excess in your buffers.

AskMe is great for a lot of things, but I'm also going to suggest that you talk to your local crystal guys about this. It sounds like you're doing a one off and sending it to a structure lab and the guys in the structure lab, if they do ANY wet lab stuff, will have a great deal of expertise. Unless YOU'RE the guy in the structure lab ... then talk to some membrane protein guys.

Good luck.
posted by sciencegeek at 4:50 AM on April 11, 2011

Also: strangerinastrainedland is absolutely correct: at a certain point, ignore your PI. Don't tell your PI you're doing a bunch of truncations, just do them and keep working on what your PI tells you at the same time, but when you're at your wits end with the full length, show him the extremely stable and hopefully crystalizable truncation/s and say, "So, we could just start with this, eh?"
posted by sciencegeek at 4:52 AM on April 11, 2011

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