Dear Beverly,

You are absolutely correct in your experience of sample loss during dialysis and buffer exchange fltration.  So, stay away from them.

Best way to remove SDS is by performing Gel Electrophoresis.  Your protein will be gel-bound, perfectly denatured and ready for in-gel digestion.  

TCA precipitation in presence of DOC (deoxycholate) is also a good way to purify your protein, free of salts and detergents.  After the precipitation step, my choice of analysis will be, again, SDS-PAGE since the precipitate will be easily soluble in sample buffer.  Just have to make sure the pH is neutral.  If the sample buffer looks yellow/green, it means that it is acidic and has traces of TCA.  Add a couple of microliters of 1M Tris to the protein solution until it turns blue.

Good luck, Hediye.

Dear Beverly,

depending on your application you could consider removing SDS before the tryptic digest by aceton precipitation. For proteins, the recovery and subsequent tryptic digest in e.g. Urea is usually very efficient (assuming you total protein content is not too low >50 ug, concentration >0.1ug/ul).

Cheers,

Daniel

Dear Beverly, 

Like Hediye, I would suggest SDS-PAGE. If you don't need to fractionate your sample, you can run a short migration in order to stack your proteins in one thick band at the top of your resolving gel (~ 5 to 8 mm thick).

Cheers,

Solenne

Dear Beverly;

I agree with all the other opinions...

and I would add  to be aware of plastic you use for further manipulation.

Other way to remove detergents from proteins is cloroform/methanol precipitation

look at: http://onlinelibrary.wiley.com/doi/10.1002/elps.201000197/pdf

It woks fine

Good luck

If there is any way you can avoid it, stay away from all precipitation and ulttrafiltration protocols out there. While it sounds great in theory, you will likely lose over 50% of your sample when doing so, or have to spend a very long time optimising all of your conditions.

The choice will depend on what proteins you are ultimately looking for. Are you looking for a specific protein or modification, or do you want to identify as many proteins as possible? If you are looking for a specific protein, then doing an SDS-PAGE is probably the best course of action. This will also purify your protein a bit and combined with a western blot, you can precisely cut out a gel band corresponding to your protein of interest.

But if you want to identify as much as possible, you will lose a lot of proteins as well during electrophoresis. Not to mention that you will need to measure 10+ samples as opposed to a single one, as you will want to cut up the entire lane. In this case, it may be best to just perform your in-solution digestion and clean-up your digest afterwards using zip-tips, for instance.

Another thing I am just wondering is why you added SDS to your sample in the first place? If it was just to aid in solubilising your proteins, you may want to replace the SDS with acid-labile detergents that will degrade after digestion when you add TFA or formic acid to your sample.

If it has some other purpose, it might be helpfull to try and find an alternative to SDS, as it should be avoided in most MS experiments.

Good luck!

Hi Beverley, I differ from Alexander. Ziptip cleanup does not get rid of SDS 100%. Intel clean up or FASP are the options. However FASP requires higher amount of starting material. In-gel cleanup  is the best way.  Also, You do not have to run the sample in a gel all the way when analyzing total protein in the sample. As Solenne already mentioned, you run the sample only into  a stacking portion on the gel  excise a thick band and digest in gel. 

Hi Beverly. My lab has tested a number of reagents and procedures for protocols that require detergent. We have optimized them on mouse lysates to provide the highest # of ions by DDA (shotgun) analysis, but I have used the same general guidelines in c elegans. I am happy to provide our standard protocols. Are you interested in a DDA (shotgun) analysis, DIA, or targeted?

1) I'm assuming your working in c elegans whole lysates? (or mitochondrial?) If that's the case can you increase your starting protein by preparing more worms? This is a relatively simple way to compensate for protein loss from cleanup procedures.

2) Is the SDS for insoluble proteins? If so, maybe try preparing the soluble and insoluble fractions separately without any detergent or cleanup procedures prior to the trypsin digestion. The trypsin digestion may still digest your insoluble proteins (maybe increase your incubation time because insolubles may be digested less efficiently). This way has been successful for me in c elegans. The downside is that its hard to know how representative BCA readings are when you have insolubles, so try to keep consistent handling of the samples.

3) If you must use detergent, try using an acid labile one like Rapigest (linked below) or PPS. If you use these, all you have to do is acidify your samples to precipitate and pellet the detergent (after trypsin).  If not those, I have heard sodium deoxycholate is friendlier than SDS but I haven't tested them directly against each other. 

4) If you still an unfriendly detergent, I highly recommend the Amicon Ultra 0.5ml 3K MWCO columns for buffer exchange. We have had great yields with those, and our collaborators have found them to work MUCH better than a number of other columns during the FASP procedure. (linked below) I have been told by some that pre-treatment of the columns and other tubes your samples go into with BSA may help with sample loss to nonspecific binding.

5) Protein precipitation always gives me significant sample losses. In comparing acetone, TCA, and methanol-chloroform methods I got the best results from MCE and TCA. MCE was maybe marginally better. 

6) We sometimes use Oasis MCX for cleaning up samples (linked). We find it gives improved MS quality and allows us to run more samples on a column before it starts degrading significantly. If you have too few samples for a 96 well format there is also single sample columns on their website. 

7) Make sure to use low-adhesion pipette tips and centrifuge tubes for anything making contact with your samples. 

Instrumentation choices and settings can make a difference as well. Someone can help if you can tell us the goal of the experiment. 

Hope this helps!

http://www.waters.com/waters/en_US/Home/nav.htm?cid=1000941&locale=en_US#FACETED_NAVIGATION

http://www.emdmillipore.com/US/en/product/Amicon-Ultra-0.5%C2%A0mL-Centrifugal-Filters-for-DNA-and-Protein-Purification-and-Concentration,MM_NF-C82301

http://www.waters.com/waters/partDetail.htm?partNumber=186000250

The simplest method I have found to get rid of SDS is to precipitate it with KCl after tryptic digest. The insoluble potassiumdodeclysulfate can be preciptated by centrifugation. SDS is almost completely removed (see paper linked below), and the KCl is removed by standard C18 solid phase extraction. I have used this method with as little as 20 µg of starting material.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3310275/

very insightful answers :)