Very Frequently Asked Questions

Since it is difficult to know the exact composition of liquid samples, proteins in solution are usually not accepted. Contact us to discuss your particular case.

Yes, this is the most important prerequisite. If you do not see a definite band by some protein detection method, you probably won't get any useful data.

The more, the better ! If you want to have a good result you should plan to load on your gel about 1 pmol of the protein you want to analyse. For a polypeptide of 50 kDa this corresponds to about 0.05 µg. It is possible to obtain results with less (down to 100 fmol starting material) but then the nature of the protein tends to play a major role. Some proteins just digest and are recovered much better than others. Remember, it is worth investing a bit more work to prepare 2-3 times more protein than having to repeat the analysis over and over.

As pure as you can get it. Remember, mass spectrometry is a physical technique, so it will detect ANY polypeptide material. Most biological samples purified from in vivo material contain relevant amounts of highly abundant cellular proteins such as actin, tubulin, or metabolic enzymes ...so you have to design a purification procedure that can resolve your (usually low abundance) protein from these ones.

No. Even if you have only one clean band by western, you probably have hundreds of proteins in your sample. Try to silver stain a 1D gel of your sample and find out if there is a band that could correspond to your protein

Run your gel, stain with Coomassie Blue, silver stain or a fluorescent dye (e.g. SYPRO ruby).
Cut under clean conditions the band you want to analyse, put it in an Eppendorf and bring it to us.

Try to work in clean conditions. The most frequent ubiquitous contaminant is keratin, that covers virtually any object used by humans. Keratin is actually one of the major components of common DUST. Wash well your gel plates. Wear gloves when loading your gels and especially when cutting the band or spot. If possible, do the cutting under a laminar air flow (hood). Use clean Eppendorf tubes.

All most common staining methods are compatible with subsequent analysis. Avoid any fixing step employing glutaraldehyde or paraformaldehyde. In general the less you fix, the better (but this is often a trade-off with staining quality). Colloidal Coomassie Blue or rapid Silver staining (see under protocols) usually give the best results.
Or, if you can afford it, the fluorescent Sypro Ruby stain

The best way to excise proteins is to use a pipette tip that has been cut so to give an aperture of 1.0-1.5 mm
in diameter and punch out a round gel piece. We both use Pasteur pipettes or plastic ones for this.
Make sure the gel is well hydrated when you cut. Cutting the gel "under water" is a good idea, it also helps reducing keratin contamination. Cutting can be a delicate procedure and you should practice on a test gel before going for a real sample.

Unfortunately protein concentration in the gel is a critical factor for the efficiency of the digestion. 0.1 ug of
protein on a 1 mm² surface will give a better yield than 0.5 ug on 20 mm². In addition, processing large gel volumes
poses a number of technical problems. Therefore, try to maximize protein concentration and only cut the "darkest" portion of your silver or Coomassie spot.

In general no. Typical cocktails for 2D-PAGE include detergents such as CHAPS or NP-40 and chaotropes such as urea and thiourea. The concentration of salts, ionic detergents such as SDS and buffers should be minimized.

We have a standard protocol for the preparation of extracts from cultured cells in our protocol page. A list of other sample preparation methods can be found on (expasy ). These protocols are a very good starting point but often need to be revised or adapted to your special case and our procedures. Please discuss with us before starting your experiments.

In an unfractionated extract of a human cell line, for example, a gel can reveal up to 3000 spots . However, these correspond often to only about 1000 proteins since many proteins generate spot trains. Extensive sample fractionation before 2D-PAGE can improve proteome coverage a lot but also generates artifacts and greatly increases the workload. Unfortunately it is a fact that proteome analysis is a lot less comprehensive than transcriptome analysis by microarray. We are only seeing the "tip of the iceberg", that is the fraction containing the most abundant proteins that can be solubilised with the used techniques.

It is difficult to answer this question. First of all the results are different and the pools of proteins analysed are not identical. To obtain characterization of more than 1000 proteins both techniques must be pushed further by prefractionation. MS has the bonus of producing a large amount of protein identity data. On the other hand, dealing with this volume of data also requires significant resources. Maybe pure MS approaches also have an advantage when sample amounts are very limiting. On the other hand, 2D-PAGE can give immediate information on post-translational modifications that may be difficult to detect by MS.