How to do intact or top-down analysis of intact proteins on an Orbitrap

It is funny that I haven't written about this before, particularly when it is such a common question for me to be asked, and even more particularly because it is so counter-intuitive.

First of all, I don't understand the physics or anything, I just have these simple concepts in my head (heck, as far as I cant tell, the physics seems a little controversial anyway).

Concept 1)  Proteins hate to be trapped.
In my head, I visualize the fact that we can't achieve a perfect vacuum, so there are some gas molecules in the traps, regardless of how well we pump them down.  The longer our big ol' proteins are in the trap, the more likely it is that they'll run into one of these stray gas molecules.

Concept 2) Crap sticks to proteins, so we need to blast them a little.
I was around when some previous students of Neil Kelleher's had a lively discussion regarding the physics around this.  My brain was it's normal reliable self and went to thinking about something like this:

Fortunately, for all intensive purposes all I really need to know is:  crap sticks to proteins, so blast them a little.

Okay, so those are my concepts.  These are directly linked to how I'm going to get a bad ass intact protein MS1 spectra:

The steps:

1) Find a nice protein standard and direct inject it.  If it is apomyoglobin, bring it up in 30% organic or higher or it won't dissolve (thanks Rosa!).  Start small, say 10-30kDa.  If doing high flow, you're going to need quite a bit.  It definitely depends on your instrument, sensitivity, etc., But for an Orbi Velos or QE, I'll probably start with something as high as 0.1ug/uL in 30-50% acetonitrile with 0.1%-0.3% formic acid.  Once I get it, I can always dilute the next injection.

2) Use the lowest resolution your instrument has.  (See, counter-intuitive, right?)

3) Fill time is not your friend, that's just more trapping time.  Keep it low, but your AGC target high (3E6 AGC, but 50 or 100ms fill time at most).

4) Microscans ARE your friend.  Rather than filling for 200ms, which is one set of proteins given a chance to react with spare gas molecules, you can do 4 microscans of 50ms, giving 4 times the number of ions 1/4 of the time to get messed up.

5) S-lens RF or tube voltage, depending on the kind of instrument, are going to be interesting things for optimization.  Mess around with them till you get the best signal

6) Adjust the spray voltage and capillary temperatures.  In general, turning them down lower than you have been using for cal mix.  These can beat up your proteins.  A lot of times if I'm using a HESI source, I just turn off the auxiliary heater (just set it to 0, it will always show you a red mark by that temperature, but that's okay!)

7) Try adding some in-source collison energy to knock some crap off your protein.  Watch for a drop in signal due to fragmentation as you raise the energy levels

8) Acquire a set number of MS1 scans.  I like 100.  Open the file, average the spectra and see how that looks.  Does it suck?  Increase your microscans and adjust all the things I mentioned above.  Try again till it looks nice

9) Are you happy with your resolution?  If no, raise the resolution, repeat steps 3-8.  If yes, move on to a bigger protein, and start at number 3 again!  Try cutting your concentration and repeating.  What is your limit of detection?  Keep in mind the rough numbers, because if you move from the ESI to micro or nano-flow, you're going to have increases in sensitivity in most (not all!  these big proteins can be harder to solvate with nano than high flow ESI).

Intacts are hard to do.  Keep that in mind.  This is a process.  It is best to start with a higher concentration of a lower molecular weight protein at low resolution and work your way up to that antibody.  Once you get a nice signal, then you can start thinking about things like SIM scans for better signal and think about fragmenting these big things!

Important note:  When you buy a protein standard, it comes all full of junk.  There are salts and detergents and preservatives and often other proteins that are in there to preserve that protein.  Most standards will benefit greatly, maybe enormously, by some sort of pre-cleanup method.