I'll give you some naive diagnosis questions I'd ask myself if i were in your place:

First, are you trying to see the mass of the whole protein? How much would that be?

Are you sure that you can detect your protein? (It would be ideal if you could check that with pure protein).

Getting a whole protein to fly well can be tricky. What about detecting it by peptide mass fingerprinting?

Also, it might be (would be strange, but might be) that the most prevalent form of your molecule was not the single-charge, so look also for (half of the mass+1)m/z.

At any rate, three things to keep an eye on for MALDI will be desalting the sample to prevent signal supression effects, trying with different matrices (I think sinapinic acid is the mot popular for protein, but there are others that could give you a good result) and trying different ways of plating (I got my best results with "half-sandwich": plating matrix, allowing it to dry and plating matrix+sample on top, but I look for a short peptide and use alpha-cyano-4-hydroxycinnamic acid, ACH, as matrix).

Good luck, and let us know how it turns out!

I'll give you some naive diagnosis questions I'd ask myself if i were in your place:

First, are you trying to see the mass of the whole protein? How much would that be?

Are you sure that you can detect your protein? (It would be ideal if you could check that with pure protein).

Getting a whole protein to fly well can be tricky. What about detecting it by peptide mass fingerprinting?

Also, it might be (would be strange, but might be) that the most prevalent form of your molecule was not the single-charge, so look also for (half of the mass+1)m/z.

At any rate, three things to keep an eye on for MALDI will be desalting the sample to prevent signal supression effects, trying with different matrices (I think sinapinic acid is the mot popular for protein, but there are others that could give you a good result) and trying different ways of plating (I got my best results with "half-sandwich": plating matrix, allowing it to dry and plating matrix+sample on top, but I look for a short peptide and use alpha-cyano-4-hydroxycinnamic acid, ACH, as matrix).

Good luck, and let us know how it turns out!

As Andres has suggested, we need more info before we can potentially troubleshoot your problem. How big is the protein, how much do you have, what buffer is it in, are you using the MS in linear mode, what matrix have you used?

Once we have more info, we can try to troubleshoot.

Peter

The protein is 25 kd fusion protein. Trx tag is 13 kd and the 12 kd the native protein. We haven't used the Factor Xa to cleave the Trx tag. Initially the protein was in the buffer containing 50 mm tris-cl, 0.5 mm EDTA, 3mm DTT and 300 mm imidazol. Then the protein was concentrated using 3KD filter in 5000 rpm. After that it was buffer exchanged with water (1 mL protein with 12 mL water) in the 3kd filter at 5000g. Then the protein solution left in the filter was used for the analysis. Before the mass spectrometric analysis I saw that it forms dimer and oligomers (SDS page band) and so I added 1mm DTT to reduce the disulfide linkage. I did not get any result in both DTT and without DTT solution. The protein concentration measured before the buffer exchange was 2.53 mg/mL, after the buffer exchange the concentration wasn't determined. Cinapanic acid was used as a matrix

If you can see the protein in a gel, it's there and should be more than enough for MALDI. You should check that you still have it, though. If the membrane of your concentration & desalting was broken for any reason, you might have lost it. It could also precipite upon buffer exchange. Let's asume it's still there. Then it's clearly a problem of getting it to fly.

First thing I'd try: different matrices. Since your protein is not very big (25KDa is for the monomer, right? Based on sequence or on electrophoretic mobility?) it might be worth to try ACHC or MBT. They work fine with peptides and small proteins.

Second: let's not discard salts and sample amount yet. When you tried different concentrations, how different they were? Did all of them have little or no salts? How much did you plate?

And last: you can always perform your Xa cleavage and see if the native protein is easier to detect (I'd go with ACHC for something close to 10KDa).

By the way, you mention that you used Cinapanic acid as matrix. I don't know anything by that name and it doesn't come up in a quick search. Is it something I don't know or maybe it is Sinapinic (also called Sinapic) acid (sounds pretty similar to me)?

Adding to Andres excellent suggestions. I always find I need to use the laser power at a much higher energy level than for peptide studies. Also, please confirm you are using the MALDI-TOF in linear mode, not reflectron mode. I also find that I need to use much more sample than what I expect. Start at as high a concentration as you can and work down from there.

I support the idea to try different matrices, it's amazing how that can affect the sample. I've attached a Sigma document that should prove helpful. Additionally, the solution you make the matrix up and how you spot it onto the target can have a huge impact on success. IN short, if you have access to an ESI instrument, I'd give that a go.

Good luck,

Peter

I don't agree that a visible band on SDS page is equal to a good result for MS analysis - in case of whole protein analysis. However, your protein concentration prior to buffer exchange sounds good.

You should check if you loose your protein on the membran, it doesn't need to be leaky for that, some proteins adsorb on surfaces.

Please check again your matrix, Cinapanic acid does not exist. Cinnamic acid is not suitable for proteins. Sinapinic acid should work. DHB might also work as it is a small protein. I like to use DHAP (Dihydroxyacetophenone) although preparation protocol is a little tricky.

I wonder about your statement that you see a lot of noise, this sounds like you are not measuring in the right mass range (and maybe not in linear mode as already has been suggested).

I remember facing the same problem like this. My problem was the sterility of my working space. You can run a gel, stain with silver nitrate, excise the band, extract the protein, and then resuspend it in a clean solution.Silver nitrate stained gel is better because it will exclude the possibilty of noise coming from the coomassie molecule. The extraction is crucial because you can also remove contaminants like sds molecule. There is a nice protocol on the web from people at EMBL, about protein sample preparation for MS analysis.

Is there any particular reason you are not digesting your protein? Detecting whole proteins is hard as you need a lot more protein to get a decent signal. Your measurement will be a lot more sensitive that way.

So, if you merely want to know if your protein is present, I highly recommend to digest your samples with trypsin first before doing your MALDI analysis. You can then use a database search (using Mascot, for instance, which is freely available on the web) to identify your protein.

another possibility is that the protein is aggregating big time into a gigantomer - such large non-covalent aggregates most often dissociate in SDS-PAGE running buffer - giving a false impression of actual size of protein under non-denaturing conditions. To confirm you have a decent size mass that your MALDI can detect, run a simple assay such as DLS (dynamic light scattering) or native PAGE.

Thanks everyone...

I used the Sinapinic acid.

I also used the reducing and nonreducing dye in the SDS page gel and I found that non reducing forms oligomers. So I added DTT (1mm in the final solution), but got the same result in the mass.

After the digestion, I got two picks----7622 and 8386 da. Whats the reason for that?

I would second Alexander's suggestion, go for an in-gel digest - this will give you the best characterization of your protein of interest. For MALDI-ToF of the intact protein, try different matrices (CHCA, sinapinic acid, ferulic acid, HABA) and also different solvents in case your protein of interest does not dissolve easily in aqeuous solvents. Isopropanol or even heptafluoropropanol with a thin-laye or sandwich preparation sometimes do the trick for sticky peptides/proteins.